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Difference between live and dead body from atomic perspective?
Sorry if this is a dumb question, recently I have been interested in the question of how "life" should be defined and got a specific question:
If we compare a live body of a person and a dead body of someone who just died, what is the difference in term of their molecule? Say for example, when the heart stops beating, the blood won't cycle through the body, but if we focus on individual cells, say the blood cell, which has hemoglobin molecules, it stops carrying oxygen when the person dies, but if we dig further, the hemoglobin molecule is defined as:

A hemoglobin molecule is made up of four polypeptide chains, two alpha chains of 141 amino acid residues each and two beta chains of 146 amino acid residues each.

A bit more further we would reach the microscopic world, where there are atoms, protons, electrons, etc, AFAIK, these things would still obey the rule of thermal motion, although the motion may be less active due to the reduced temperature of the dead body.
So from the atom's perspective, what is the difference between a live person and a dead body?
Thanks and sorry again for this dumb question!

From atoms prospective there is no difference. It is like a car that has crashed: from the atomic (and even material) perspective it is still a pile of metal, glass, and plastic... but the shape of the parts and their position in respect to each other are such that it cannot function anymore as a car. One could say that being a car is more than just being a collection of atoms - it is an emergent property. You can search this site for emergence.

Thanks @RogerVadim to me the car example feels a bit different, a dead body can be lifeless without any damage or deformation, yet there is no life in it, the atoms, molecules, organs, skins, etc, are still in good shape, it's more like a machine that runs our of power and no power can be provided to it for some reason, and comparing such a machine with a running machine that does have power supply, there's no difference even between parts / components.

@hzhu What you say is correct. The difference is that car materials are stable, soone can restart it long time after it stopped, whereas the molecules in an organism start to degrade. If new molecules are not synthesized, this degradation becomes irreversible. In other words, there is damage, even though it is not immediately visible. Although forensic scientists are trained to see even insignificant markers.

None.
Say you were to take a skin sample from a living individual, and a skin sample from a deceased person. If you were then to somehow extract and isolate a single atom from each sample, they would be indistinguishable. In other words, if you were to mix up the two atoms, there would be no way if telling which came from the deceased individual.

From an atomic level which I believe you are after, the difference is quite simply, the living body has electricity running through it.  Without electricity the atoms in your body could not be manipulated by cells to become more negatively and positively charged leading to a voltage and ultimately a current the central nervous system takes as a signal for your heart to beat, for an arm to move, to breath.  So atomically the living and the dead are the same.  But the living person through the process of the nerves and other cells create electricity through a process of letting in sodium or potassium.  Obviously food is where we get the atoms to not only build tissue but the energy to make our own electricity.


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The very first living organism model
Can you advice me where I can find explanation, what was the very first living organism on Earth? I am looking for nicely explained material, preferably with some 3d models and animation, showing how it could work. Maybe a computer emulation?
The sources I was able to find so far were very theoretical and dry, they didn't allow me to get a good grasp on the concept. Basically I am looking for the simplest living organism possible in order to get a good grasp on the mechanics of life.
From the sources I studied so far I understood that it was just a molecule of RNA, which could copy itself, right? But I didn't find exact description how it could look like..

Not the first living organism, but Julia Child on how to make primordial soup: video

Nobody knows for sure -- there are many theories, tending to be quite general. "RNA world" is indeed one theory, but there are others. You will become quite celebrated and famous if you end up providing an answer for your question.

Does this answer your question? Thermodynamically, how did the first cell arise?

@Armand Haha, yes! After asking the question I found out that nobody got an idea what was the earliest self-replicating structure in the first place. And I wanted to see a nice simple model of it!

@Luigi Not quite, but it is an interesting information. I wanted a visual easy to understand model of the first structure that could replicate itself. But it seems that it is yet unknown.

Fundamemtal basis of establishing something as living is the capability of replication and producing copies of forms similar to itself. Therefore, something which has two properties  i.e to act as enzyme and information carrier at the same time can perform that function and qualify to be called as first living *(but we will call it first living system or molecule not first living organism some say so, there are many arguments, depending upon how they look upon something to be called as living, it is itself a big question but it seems strongly persuasive that something like RNA world model is surely first form of life) This system is self perpetuating(heredity is basis of evolution and this is basically  heredity). If you want the story behind how it was done and quench your thirst (its short read) I'm giving the link below
https://www.sciencedaily.com/releases/2009/01/090109173205.htm
After you'll read that It was not easy to make this cross replicating system. And get that result to establish grounds of RNA world.

source: Stack Exchange

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

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How RGB cones see other frequencies?
Orange,violet and yellow have different frequencies.
How can we see them, as we have only three receptors rgb?
Merely combining those frequencies cannot generate a new one.

This question has rather detailed answers in the physics community. In a nutshell - they don't see other frequencies, but rather the color is determined by our brain by the relative excitation of different cones.

How does your compute display create all sorts of colors when it has only 3 colors of pixels?

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


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can a cell evolve into a different species in the lab?
A friend of mine does not believe in evolution. He claimed that we can not as humans observe a single cell evolving into a different cell. Is that possible to be observed in the lab?
Thanks in advance.

First it is worth noting individuals don't evolve, populations evolve.
We can observe cells giving rise to different cells, the long term E.coli experiment is a famous and ongoing example, not only can evolution be observed but because sample cells from the population are taken and frozen periodically, it can be repeated! that is you can observe a change then set up the same line of cells under the same or under different conditions to see if the changes are different. In some cases even the exact same mutations can be observed.
https://en.wikipedia.org/wiki/E._coli_long-term_evolution_experiment
http://myxo.css.msu.edu/ecoli/
https://www.pnas.org/content/105/23/7899

+1 Moreover, building devices to observe evolutionary changes has become nearly "popular mechanics" level activity: morbidostat


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Can one identify an organism based on its genome alone?
If one is given the complete genome of some unknown organism, would it be possible to systematically deduce what this organism looks like and behaves like without reference to anything else (ex: a repository of genomes of known living organisms)?
In another words, do we currently have a model capable of translating any random genome of an organism to a graphical representation of the organism and information on its behaviour?
Are there such 'genome-structure' or 'genome-behaviour' relationships known to us that would enable us to accomplish such a task?

As you specifically state:

Would it be possible to systematically deduce what this organism looks like and behaves like without reference to anything else (ex: a repository of genomes of known living organisms)?

No, no chance at all, we certainly wouldn't even be able to determine a single gene function, other than that it codes for a series of amino-acids. After all, even simple genes of say 100 bases in DNA will still have 33 amino-acids, and most people don't remember that sort of information without cause, and the only way we can acquire that easily information is through repositories of information. The only way we could do something approaching working this out would be to take portions of the genome that we could identify as genes (start/stop codons), then express them one-by-one and empirically work out the function through classic biochemical methods. This is very laborious and time consuming - it would take many many person-hours per gene and a LOT of resources. However, this is how functions of novel genes and their related proteins were and still are worked out.
With reference - perhaps, at least partially. From the sequence and with access to a repository such as Genbank we could deduce which type of organism it was by comparing to known genomes from other organisms in a process known as phylogenetic analysis. You could certainly tell which of the kingdoms it came from. In each organism there are conserved bits of the DNA (or RNA) that tell you which groups they are. For example all DNA based bacteria and archaea (that I know of) you can use the 16S ribosomal RNA sequence, which will identify bacteria and archaea down to genus and species level.
For instance, you might have a genome that gives you information that it is a bacterium of the Staphylcoccus group, meaning that you can deduce that it is very very likely that it will be a small bacterium which is a round ball shape, and will stain positive with a Gram stain. You could also likely identify some genes and their function based on homology.
You can apply this process to other genomes, for instance a mammalian one will tell you that it had fur, produced milk, was warm blooded, quadruped, spinal column etc.

No, current technology is nowhere near what is required to deduce the form of a species from its DNA alone without comparing it to the DNA of the same or similar species.
The reason for this is that genes in the DNA generate proteins and these proteins are simple building blocks that combine in highly complex ways that we do not understand. Currently scientists are simply trying to learn how to predict the SHAPE of the protein from its DNA sequence. That is the first step. We are getting close to being able to do that. Once we can predict shape, then we can start working on figuring how that shape translates into function.
I would also add that organisms have many expressed traits. That means that the trait expresses itself only after the organism grows and is not genetically determined. For example, fingerprints are an expressed trait. Also, colorings and pigmentation are expressed traits. For example, I have two cats that are genetic twins and have exactly the same same DNA, but one has a small patch of white fur on his chest and the other does not.

However, there are individual genetic variants with known function. For example, given a person's genome, we should be able to roughly predict what hair color and texture they have, eye color, etc.

@Armand, that doesn't fit the question, which specifies "without reference to anything else (ex: a repository of genomes of known living organisms)".

@Armand we know only that specific genes correlate with specific hair colors, but we don't know the full cause-and-effect chain. In the end, to determine "this gene encodes red hair" from first principles, you'd have to know the function of all the genes that are involved in creating hair cells, pigments in general, and how the gene is activated in hair cells but not in skin cells.

Yes, by OPs mentioning a "model" I was assuming that model would contain info like hair color genes and so forth. In re-reading the question it does seem the OP intends a de novo model based on structure prediction.

Echoing other answers, our ability to predict the function of individual genes is entirely dependent on either A) physical experiments (making the products of those genes in a test tube or performing genetic experiments) or B) inferring their function from their similarity to other genes that have been studied by physical experiments.  In other words, we can't currently predict individual gene functions with computers unless we have some similar outside reference to compare to.  Without the ability to do that for a single gene, there is no possibility of doing this for a whole genome.  Forget about predicting how all of these genes would express or interact to create a complex living organism.  If you could compare to other sequenced genomes, phylogenetic analysis would quickly tell you what species your unknown genome was most similar to.
I thought I might offer an alternate answer about what one might be able to do with some general knowledge of genomes but not the ability to actually compare your unknown genome to specific genomes / known DNA sequence.  With the right software, you could roughly predict where the genes were in your unknown genome.  This could tell you how many genes this organism has, how many of those genes are unique genes or repeats, and how much of the genome is made of non-genic sequence.  You could also look at the overall structure of your genome.  These factors could give you a rough guess about what kind of organism you have.  For example, if your genome is circular (which you could tell from the sequence), you have a prokaryote or archaeon.  If there are very few genes, you could infer that this organism lived a parasitic lifestyle, since the smallest known genomes are from parasitic species that no longer need to encode for all of the functions of a fully independent lifestyle.  A large genome, linear chromosomes, or multiple chromosomes would indicate a eukaryote.  If you see evidence of polyploidy, your organism could be a plant.  Through similar methods, you could probably tell a eukaryote's nuclear genome apart from mitochondrial and chloroplast genomes and guess whether it was a plant based on how many non-nuclear chromosomes it has (two distinct non-nuclear chromosomes would indicate a chloroplast genome is present).  Just a little thought experiment on what one could do with an unknown genome sequence without Genbank or reference sequence, but with current computational tools and some knowledge about genomes.

How would you know (without reference to genbank or similar) that a particular gene came from a mitochondria or a chloroplast? I can see the bit about eukaryote vs prokaryote, but I don't see that the next steps would work. I don't think people have solved how to determine ploidy with sequencing yet - please let me know if I am wrong, because I love to learn new methods.

Ploidy is a number-of-copies measurement.  If you had error-free sequence for all the DNA from a single cell, multiple alleles for the same gene could be tallied separately. You might reasonably infer triploidy if you found many genes with three alleles present, tetraploidy if you found many with four alleles present, etc.   Of course you wouldn't expect multiple alleles at every site,  but finding even a few such would be a strong hint.

@bob1, for mitochondria or chloroplast chromosomes, they're both circular (although wikipedia tells me that there's some debate about chloroplasts).   So, they should be identifiable apart from other eukaryotic chromosomes without having to worry about identifying the genes themselves.  Also, I'm guessing that both mitochondria and chloroplast chromosomes are much more gene-rich than most nuclear chromosomes, so they could probably be confirmed with that as well.

@bob1, as to polyploidy, I had allopolyploids in mind like wheat and other plants, which I think would be strikingly obvious if you were to look carefully at your sequence.  I'm not experienced in this area at all, but Ethans answer sounds pretty logical to me for cases like autopolyploidy.  That would depend on having a fair amount of heterozygous alleles though.

@Ethan - but modern sequencing technology (with perhaps the exception of PacBio and Oxford Nanopore) gives varying sequencing coverage and depth for a single sample, which in no way relates to ploidy as far as I can tell. There are tools out there to work this sort of thing out, but as far as I can tell they are  definitely imperfect to put it mildly.

@LTRretro Yes, mito and chloroplast are classically described as circular - my question here (and I should have been more clear) relates to how you would tell the mitochondrial/plastid genes from those of contaminating DNA from bacteria etc. These generally make up a significant portion of any sample taken from an animal, with the exception of deep tissue, even then you tend to get some.  But I guess this is getting away from the topic in hand, where one is given a complete sequence, not just the raw reads.

@Bob1 - The idea is not to rely on quantitative estimates of copy number.  The idea is to look for loci where N sequence variants are present.   In practice most of these will be artifacts, but as the hypothetical error rate goes to zero any remaining cases are due to multiple copies of the gene.  That would include paralogs and multi-copy repeats within a single genome, but also it would include the presence of multiple alleles due to polyploidy.


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Can we improve the hemocompatibility of VAD by growing a single layer of endothelial cells on the surface?
Maglev centrifugal pumps have been used in the latest ventricular assist devices (VAD) and several experimental total artificial hearts (TAH). Compared to previous generation pumps, maglev pumps don’t have any moving parts (joints, bearings, etc.), which greatly reduces the risk of coagulation. However, the risk of blood clotting is still not eliminated. Normally the bloodstream is lined with vascular endothelial cells, which secret anticoagulants to prevent blood clotting in the vessels.
So I come up with an interesting idea. Can we prevent blood clotting by growing a single layer of endothelial cells on the surface of the stator and rotor? We have successfully cultured many animal cells. By adding a few animal cells and culture medium into a bottle, the cells will grow into a single layer of cells along the smooth surface of the bottle. Because there is no moving part inside maglev pumps, it would not be difficult for endothelial cells to adhere onto the surfaces. The blood can serve as an ideal culture medium which provides nutrition and carries away metabolic wastes. So if we can grow endothelial cells on VAD or TAH, will it eliminate the need to take anticoagulants?

"By adding a few animal cells and culture medium into a bottle, the cells will grow into a single layer of cells along the smooth surface of the bottle" - oh really? What makes you think that?

So what else does it take?

The answer to that question would fill textbooks.

I know there are many other things required like temperature, pH, disinfectants and growth factors, but I don’t think this is a big problem inside our own body.

Our body is the best cell culture.


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Can genetic recombination occur between DNA viruses?
Single stranded RNA viruses like coronaviruses can recombine with each other. During the viral RNA replication, the viral polymerase can jump onto another template strand and result in a chimeric genomic RNA.

However, trying to extrapolate this model to DNA viruses will encounter a big problem. Most DNA viruses have double stranded genomes. The replication of dsDNA is semiconservative, which means the template (parent DNA) is also a part of the daughter DNA. Even if the DNA polymerase jumps in the halfway, the template DNA won’t move with it. Does that mean DNA viruses don’t recombine, or recombine in different ways?

Summary
Certainly DNA viruses do recombine — some of them encode proteins specifically for this purpose. And this is unsurprising if one reflects on the fact that recombination is a fundamental characteristic of double-stranded genomic DNA in prokaryotes and eukaryotes. What is surprising is that recombination occurs in RNA viruses — the starting point for the question. However the mechanism by which this can occur — apparently in a random manner — is completely different from the long established way in which double-stranded DNA genomes recombine.
Apologia
The literature on recombination is vast. For the purposes of an answer to this rather naive question, I think it sufficient to refer the reader to a Wikipedia article on recombination in general, and draw on a single review of recombination in DNA viruses.
Answer
Quoting from the aforementioned review by Weller and Sawitzke in Ann. Rev. Microbiology (2014) 68, pp 237–258:

“Homologous recombination plays many important roles in the biology of all living organisms including DNA replication and repair of DNA damage. Recombination is essential for genetic diversification required to enable organisms to adapt and evolve.“

The Wikipedia article deals first with ‘standard’ recombination in DNA genomes, which, although not identical in all situations, generally involves a double-strand break, pairing to a homologous region on another chromosome, and often ‘crossing-over” of the strands between the two chromosomes:

This is illustrated in more detail in the article, but one should emphasize that crossing-over involves an intermediate Holliday junction of four strands of DNA, as shown below for PDB file 1M6G.

The idea that single-stranded nucleic acid is a requirement for the process (as assumed by the poster) is false.
Recombination is catalysed/promoted  by specific proteins — recombinases — the most well-known being perhaps RecA in Escherichia coli and Rad5 in eukaryotes. Specific nucleases are also involved in making the strand breaks. What the review by Weller and Sawitzke focuses on is the evidence for similar recombination and recombinases (and nucleases) in DNA viruses as diverse as bacteriophage lambda and herpes simplex virus type 1. They examine the mechanism and identify the viral and host proteins involved and conclude that:

“Both bacteriophage λ and herpes simplex virus display high rates of recombination, both utilizing their own proteins and commandeering cellular proteins to promote recombination reactions.”


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Many viruses use pressure to pump viral DNA into host cells. Does that mean these viruses have a narrow acceptance of the genome size?
DNA viruses like bacteriophages and herpes viruses have rigid capsids. The viral DNA is densely packed inside, and the internal pressure is 1~2 orders of magnitude higher than the atmospheric pressure. When the capsids open, the viral DNA is pumped out by the pressure. Considering that DNA is not easily compressible like gas, a slight variation of its length will result in dramatic change in the pressure. As a result, if the DNA is slightly longer, it would be difficult to package it into the capsid during the assembly. If the DNA is slightly shorter, the pressure may be too low to pump the viral DNA out during the cell entry.

Please quote a source for these claims, as the idea of pressure within viral capsids is new to me at least. How is this supposed pressure generated and what form does it take? Are you saying that there is compressed air there? Are you aware that two orders of magnitude means “100s”?

Stress in the capsid is the dominant factor throughout early ejection, with the pull due to binding particles taking over at later stages.
https://www.sciencedirect.com/science/article/pii/S0006349506717428

There isn't enough data to give a clear answer to this question, but I can at least explain what I know.
First, it's not 100% certain that genome ejection of tailed phage and herpesviruses depends on stored pressure in the capsid. This is called the continuum mechanics model; but another possibility, the hydrodynamic model, suggests that the difference in osmotic pressure between the cytoplasm and outside environment allows water to flow through the capsid and tail into the the cell, dragging the DNA payload with it. Personally, I am skeptical of this second model because it doesn't seem to explain herpesvirus genome ejection, and it's not consistent with the lethality of R- and F-type pyocins, which are essentially just decapitated phage tails. But there is still some question as to how the genome gets fully out of the capsid.
Assuming that the continuum mechanics model is closest to correct, the nature of the pressure generated when DNA is loaded into the capsid by the packaging ATPases is not intuitive. From a 2013 review:

At this concentration, DNA is highly condensed, and much of the water that normally hydrates the DNA and its counterions must be removed in order to package a complete genome. Water removal during the packaging process occurs by reverse osmosis, the energy for which is derived from the activity of terminase.

Only about 10% of the energy stored in the packaged phage DNA is due to bending DNA more tightly than its persistence length. Most of the energy internal to the capsid is due to the dehydrated state of the DNA, which is reflected by internal osmotic pressures that reach tens of atmospheres.

...And from the herpesvirus paper linked above:

DNA can fill as much as 60% of the internal capsid volume, with the remaining volume fraction occupied by water molecules and small ions that freely diffuse through the capsid wall.

So, despite what we'd expect from our macro-scale experiences, the DNA IS compressible almost like a gas. Small differences in the length packaged won't change the final pressure dramatically.
Also, many of these viruses use a so-called "headfull" packaging method. The viral genome is copied continuously, with the end of one copy linked directly to the beginning of the next copy. The ATPase motor packs this continuous chain of genomes into the capsid until the pressure reaches a certain level, then the motor stops packaging and cuts the DNA chain into a piece inside and a piece outside.

When a threshold amount of DNA, representing
about 103% of the SPP1 genome, has been packaged (headful) a
sequence-independent DNA cleavage terminates the encapsida-
tion cycle (Fig. 1A).

So what will happen to a variant of the phage mentioned here, SPP1, if an insertion mutation increases the length of its genome by more than 3%? I don't know, but in my experience with capsid assembly, it's not uncommon for a small number of large capsids to be produced along with the normal capsids, so possibly large-genome mutants can get by until the structural proteins co-evolve to make the larger capsids the new genome requires. Alternately, even if individual virions no longer contain the whole genome, if multiple virions infect a single cell, all genes will be present in at least one copy, again giving the structural genes time to evolve.

That makes sense. If DNA is very stiff like water, the pressure will lose quickly as long as only a little DNA is spit out. So DNA must have an effective “bulk modulus” like gas, which enables the capsid to pump out the DNA continuously. Whether the pressure is due to compression, dehydration or Coulomb repulsion is insignificant.

The mechanisms that generate pressure inside the capsid are complex and nonlinear, so you can't really assign a "bulk modulus" to the whole thing. You could probably estimate a modulus for when capsid is 0-30% full, another for 40-55% and another for >60%, with intermediate values in between where no one mechanism is dominant. All the phage motors I know of can "switch gears" to package rapidly when there is low resistance, then package more slowly when resistance is high.


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Do endogenous retroviruses (ERV) have a high degree of heterogeneity among people?
Endogenous retroviruses (ERV) make up 5~8% of human genome. These ERV are ancient retroviruses which infected germ cells and therefore became parts of our genomes. However, unless the ERV can increase the survival chance to its host, it wouldn’t be able to expand its frequency in our genetic pool. So there must be some mechanisms that expand the copy number of ERV. The infection of germ cells can happen more than once. Alternatively, ERV can reactivate at some point of the development of the host and make new copies in the genome. Considering that the sites of integration of retroviruses are highly random, even the same ERV can appear in different loci or different chromosomes. Have large scale genome sequencing revealed a heterogenic map of ERV?

Of the papers I reviewed to answer this,
Variation in proviral content among human genomes mediated by LTR recombination
addresses the question most directly. I recommend you read it if you have access. To summarize, most ERV in the human genome integrated before the separation of human and chimpanzee lineages, and some before the emergence of placental mammals. Among these, there is relatively little variation:

an HERV-H element on chromosome 1 (1q25.3_H3) was shown to exist as proviral and solo LTR alleles in two related individuals

"solo LTR" allele means that the viral genes have been deleted by homologous recombination of the Long Terminal Repeats (LTR) that flank the intact provirus, leaving only one LTR. These don't code for any protein, but can still be "active" by affecting the expression of nearby regions in the host genome. The paper I cited above verified four other HERV-H variations and identified 67 potential variations.

We were intrigued to discover a dimorphic element for the HERV-W family (18q21.1_W2). This element is represented as a solo LTR in the reference genome, but our data clearly show that it also occurs as a provirus segregating in South Asian populations (Fig. 3a) and likely in other diverse populations (our pipeline predicted a provirus allele in 194 out of 279 individuals surveyed, Additional file 2). To the best of our knowledge, this is the first HERV-W locus reported to show any type of dimorphism. This particular HERV-W insertion must have occurred between 18 and 25 million years ago because a provirus is found at orthologous position in all other ape genomes including gibbon, but is absent in Old and New World monkeys [67]. Our discovery illustrates the potential of LTR recombination to alter genome structure long after a proviral insertion has occurred.

The HERV-K family, however, integrated into the human germline relatively recently (estimates span 3 million - 250 thousand years ago).

Among the diverse assemblage of HERV families in our genome, a single subfamily known as HERV-K(HML2) has been reported to exhibit insertional polymorphism in humans [17, 28, 29, 36–47]. Thus far, approximately 50 HERV-K(HML2) proviral loci are known to occur as empty (pre-integration) and/or solo LTR alleles segregating in the human population [17, 43, 45, 46], but more may be expected to segregate at low frequency [39, 48]. These observations are consistent with the notion that HERV-K(HML2) is the most recently active HERV subfamily in the human genome [49–53].

The 2020 review article Human Endogenous Retrovirus K (HML-2) in Health and Disease gives a quantified overview of the variability of the HERV-K family.

It is known that there are more than 1,000 HERV-K (HML-2) loci in the human genome. Most of them reside in the genome as solo-LTRs generated by homologous recombination between the LTRs of a single HERV-K (HML-2), resulting in the deletion of the internal sequence (Hughes and Coffin, 2004). Solo-LTRs are approximately 10-fold more abundant than their full-length or nearly full-length proviral integrations (Subramanian et al., 2011). In total, there are 1098 reported loci whose insertional elements possess functional elements, including 1061 reference (hg19) insertions and 37 non-reference insertions (Subramanian et al., 2011; Lee et al., 2012; Marchi et al., 2014; Wildschutte et al., 2016; Wallace et al., 2018; Xue et al., 2020).

My summary: Most of the ERVs in the human genome are present only as remnant solo LTR alleles. Since virus reemergence is harmful, deletion by homologous recombination is likely selected for. This kind of deletion has been happening continuously along the human lineage and accounts for most of HERV differences between human genomes. Only the HERV-K family shows differences in insertions between humans. Across humans, roughly 95% of HERVs are identical for everyone, and roughly 5% are absent in some people. All of the HERV families are considered extinct. That is, there are no retroviruses circulating currently that correspond to any of the integrated HERVs.
I didn't see much discussion about smaller-scale insertions, deletions, or point mutations within HERVs, except to note that intact provirus could be dated by the amount of sequence difference in the LTRs at either end. That is, the LTRs are by definition identical when the retrovirus genome is inserted, but will accumulate mutations independently afterward, so the difference lets you estimate the number of mutations and thus how long ago integration occurred. If the LTRs of a provirus diverge enough, they will no longer be able to homologously recombine, and the virus insertion will become permanent. As a caveat, genomics is not my specialty; I may have combined information from different sources incorrectly.
some other notable references:

Identification of the distribution of human endogenous retroviruses
K (HML-2) by PCR-based target enrichment sequencing
Is the
HERV-K HML-2 Xq21.33, an endogenous retrovirus mutated by gene
conversion of chromosome X in a subset of African populations,
associated with human breast cancer?


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Can Covid's RNA be nullified by an opposite RNA strand?
Firstly, I have scant knowledge of biology, epidemiology, virology, and related fields. If my question seems nonsensical or bewildering, I beg your pardon. I'm not sure how to phrase this, but I'll try my best.
As far as I know, when the Covid virus enters your cells, it releases its RNA so that it can copy itself and replicate and thus spread in body.
Can that Covid RNA strand be "nullified" when it comes in contact with an opposite RNA sequence? In other words, just like if you pour water on a fire, the fire will extinguish and thus the two elements can be considered "opposites" as per my logic. In the same way, can an "opposite" RNA thing exist, which can possibly "extinguish" the Covid RNA? If you know what I mean. The analogy is that the "opposite" RNA thing is water while Covid RNA is the fire. The former can take out the latter.
Is there something that works like this? If not, is it possible to develop something that works like this?
Because that way, the Covid replication process is halted at that first step, and ergo the virus can't infect the body.
Thanks.

Yes is the short answer.
These are known as silencing RNAs or interfering RNAs
The longer answer is
You would need much more RNA than you could easily administer to the epithelial cells of the mucosa and other tissues that viruses like SARS-CoV-2 infect, so as to cover each individual cell. In addition you would need to be able to deliver it to those tissues in a timely manner (inhalation is possible, several drugs are administered in that manner), have it enter the cells (essentially what the Pfizer/Moderna vaccine does)  and not have it be degraded.
These are not insurmountable problems, the Moderna team managed to do it for their much longer vaccine, but I don't know if the same base modifications that work for their vaccine will still work for interference/silencing.

I suspect that delivering to all the cells infected by the virus in large enough amounts to slow viral replication is indeed an insurmountable problem (otherwise the virus would have unprotected cells within which to replicate); the Moderna vaccine just has to get into some cells in modest amounts, not all affected cells in overwhelming amounts.

@Armand - I totally agree, but it could be possible to do, just extremely impractical and probably with horrific side-effects, if the transfections that I do regularly, can be taken as a basis.

Hmm silencing siRNAs notwithstanding I’m not happy that you’re answering OP’s question with “yes” since, frankly, the concept of “nullifying” or negating RNA doesn’t really make sense and does certainly not exist: the RNA is neither “extinguished” as in OP’s analogy, nor otherwise negated or nullified. No such effect exists: opposites don’t cancel out (RNAs are not physical forces), and what would this even mean for RNA?  The fact that RNAi superficially looks like it does exactly that is all the more astonishing but unlike OP’s proposal there’s actually a logical mechanism underlying it.

@KonradRudolph - if you can make a ssRNA into dsRNA the viral RdRP will no longer work on it properly, especially if you can make some of the bases non-conventional.


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how to perform ligand and DNA alignment in pymol
I need help in inserting ligand to specific base pair in DNA. I have a drug molecule and want to insert in between the base pairs I am interested in (it is dsDNA). Please note that it is a customed sequence dsDNA so it does not have any intercalation site for the ligand to bind itself. I tried to follow the tutorials but cannot find something where I can insert ligand into the custom DNA sequence I have generated.
I will appreciate if you can please help me or guide me where I can study and understand how to do it.

Welcome to Biology.SE. Is there some reason you are asking this here rather than in [Bioinformatics.SE]? If you agree that site seems more appropriate please either delete from this site and repost there or request migration. Please do not crosspost as that is considered to be impolite.

Could you clarify? Am I right in thinking that you are trying to answer the hypothetical question: “If ligand L were to insert itself between two specific base pairs in my custom dsDNA structure, what molecular interactions would it make?” If so, have you any reason to  think an application to display molecular structures can do this? I would except to have to use some high-powered energy minimizing program to compute possible structures first. Are you sure Pymol has such a facility?


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What is the probability distribution of a neutral mutation's allele frequency after k-generations? (in asexually reproducing organisms)
I've had trouble finding info on how to model this problem, so figured I'd ask it here. I'm trying to figure out what the probability distribution is of the frequency of a mutation in a population with no affect on fitness as k generations after occurring (assuming stable population size N). Obviously there are other parameters such as lifespan, but the simplest version would be that each generation, every cell (it's a single cell organism) either undergoes cell division, leaving two daughter cells, or it dies, with probability 0.5.
E.g., a mutation that occurs at t = 0  has probability 0.5 of going extinct and the same probability of being present in 2 cells (thus frequency x = 2/N) after one generation; then at t=2 it p(x = 0) = 62.5; p(x=2/N) = 0.25; p(x=4) = 0.25. How do I generalize this to get a distribution for after k generations? Thanks.

I think that you are interested in the original work about neutral mutations from Kimura (1954, 1962). This work established that the probability of fixation of a neutral mutation is equal to its current allele frequency.
The take-home message is that the neutral mutations are taking a random walk through allele frequency space, i.e. a diffusion process, which makes it tough to come up with well-behaved distributions.
You can model this with the normal Wright-Fisher model in the absence of selection (page 19 here). It's a bit of a headache because it involves following a Markov chain for $k$ generations. Possibly the most straightforward thing to do is to just simulate a million trajectories using the Wright-Fisher model and see what they look like for any given $k$ and a new mutation. For an example of how to do this look here, or alternatively you could use an existing program like SLiM.
If you are looking for something a bit more closed form, what Kimura (1954) did was to apply a diffusion model approximation of the Wright-Fisher model (equation 1), with some further development.
He writes:

If $\phi(x,t)dx$ is the probability that the gene frequency lies between x and x + dx in the t-th generation, it can be proved that $\phi(x,t)$ satisfies the partial differential equation,
$\frac{\partial \phi (x,t)}{\partial t} = \frac{\partial ^2}{\partial x^2} [\frac{V_{\delta\_{x}}}{2} \phi(x,t)] - \frac{\partial}{\partial x}[M_{\delta_{x}} \phi(x,t)]$

This is where my math fails me in population genetics and I am less able to help you applying and untangling how to come up with a distribution. But I would suggest reading through that Kimura paper and using it as a starting point to do some research and see who cites it to give some kind of solution that's more useful for you.
A treatment of this in slides for a class is given here.


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What is the mathematical relationship between selection coefficient and dN/dS
dN/dS is often used as a measure of the intensity of selective pressure on a mutation or gene, but I'm curious about how it can be written as a function of the selection coefficient. I'm specifically interested in the scenario of somatic mutations in human cells; suppose there's a gene, and all non-silent mutations in the gene are equivalent (such as if they're all loss of function mutations). The mutation rate is μ mutations per bp per cell division and once both alleles are (nonsilently) mutated, there's no fitness gain (or loss) from additional mutations. The selection coefficient for mutations in the gene for homozygous mutations is:
w_hom = 1 + s

and for heterozygous:
w_het = 1 + hs

How would one derive the expected dN/dS ratio (where the selection coefficient of a silent mutation is 0)? I realize there are a few questions for which answers must be assumed to begin to answer this question, such as:

Is population static, or growing?
Is dN/dS only including clonal/fixed mutations, or perhaps all mutations with a frequency in the population greater than or equal to some value?

I'd be interested in what the relationship with dN/dS is even under the simplest assumptions, e.g. h = 1, only considering clonal mutations, and static population. I've found one paper that posits such a formula, citing a finding from another paper, but I don't think either shows all the math.

Are you suggesting that all non-synonimous mutations have the same fitness? Also, I think homo vs. heterozygocity is not really the issue here, unless we also need to account for recombination.

All non-synonymous mutations might have the same marginal effect fitness, in that they all just break the copy that gets mutated, but having a cell with one remaining functional copy may be more/less fit than one with no functional copies remaining.

Now you seem to suggest that these mutations are wekly deleterious. Perhaps, you could add a reference to what you consider standard use if dn/ds - we might be talking about different things.

dN/dS is usually calculated for an alignment or a phylogenetic tree as if the gene is haploid. Dioloidy is an interesting complication, since it can render deleterious mutations neutral. Selection coefficient can be calculated, if we make additional assimptions about the population growth, etc., as suggested in the OP. However, ut would limit the applicability of the method to the situations covered by these assumptions.

References
Perhaps this article is relevant, as it associates the fitness edge (i.e. accumulation of multiple beneficial mutations) with dN/dS:
Predicting evolution from the shape of genealogical trees

I think an example of it being used in this kind of context is in this paper: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC5720395/. They're using dN/dS to assess selective advantage/disadvantage of mutations in cancer.


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What exactly does my body do with oxygen? Why do I need it?
I know that my body and brain need oxygen in order to survive. If I stop breathing, my lungs (and blood) can't distribute oxygen to my body anymore, and I could get brain damage and die. What exactly is being done to the oxygen, and why do I need it?

it is expected that you atleast do some research before asking a question here,this is a starting point for research into this subject https://www.nursingtimes.net/clinical-archive/respiratory-clinical-archive/every-breath-you-take-the-process-of-breathing-explained-08-01-2018/

Oxygen is used to oxidize (burn) molecules and generate energy.
A fire combines oxygen with carbon containing molecules (like coal) and releases energy (heat) as well as carbon dioxide and water.  A fire needs oxygen or it will stop.
You need oxygen because inside your cells is an oxidation process much like a fire.  Oxygen is combined with carbon containing molecules like sugar.  When they combine, energy is released.  As opposed to a fire which releases the energy as heat, your body captures the energy in a chemical called ATP that can then be used in the body to do a little dance, and make a little love.
The oxidation reaction in your body also releases CO2 and H2O which you exhale.


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Are the terms "provirus" and "prophage" interchangeable?
Regarding the difference between proviruses and prophages, Wikipedia states that "these terms should not be used interchangeably. Unlike prophages, proviruses do not excise themselves from the host genome when the host cell is stressed." However, Khan Academy implies at 3:41 that the two terms are interchangeable. Who is correct?

Over some 15 years in academia and research, I've taken graduate-level virology classes, worked in multiple phage research labs, and attended dozens of seminars on both bacterial and eukaryotic viruses. I can't recall ever seeing or hearing the term "provirus" used until reading this post.
Sometimes different disciplines in biology will use the same or similar terms somewhat differently. There are, in fact, several examples of commonly used terms in biology that have no universally agreed upon definition. However, according to at least one source that I just found on a plant viruses, a "Provirus is a permanent part of host genome," meaning it is not excised before reproduction. This other source uses provirus to describe integrated HIV, which also doesn't need to be excised in order to reproduce. So there may be some consistency of usage here.
Ultimately, meaning in language comes from the intersection of use and understanding in context. Any time you're talking about viruses that infect bacteria and/or archaea, the terms "virus" and "viral" can almost always be replaced with "phage", but don't always have to be, so could be somewhat interchangeable in that context. Whereas, to my knowledge, "phage" is never used in describing eukaryotic viruses.
So, if you said "provirus" in a phage biology lecture, your audience would likely understand it to have the same meaning as prophage, but if you said "prophage" to describe an integrated human retrovirus or retroelement, you'll almost certainly confuse a part of your audience, so not really interchangeable at all in that context.

Since phage are bacterial viruses, every prophage is a provirus, but not all proviruses are prophages. The term "provirus" IS a thing in molecular biology, but as you note is probably much more common in work on retroviruses or eukaryotic genome evolution.

@Armand I agree about prophages being proviruses. One difference being that lysogenic prophages are excised from the host genome before a lytic replication cycle can begin, where retroviruses can generate genomic viral RNA (and infectious virions) while the provirus remains integrated on the host genome. So IF being a PERMENANT part of the host genome was a necessary property of all proviruses, then prophages would not technically be proviruses. But that would seem like a pretty arbitrary distinction to make (imho).


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If the AraC protein was a gene repressor when it binds arabinose, would there be high or low transcription levels when arabinose is present?
If the AraC protein were to function as a gene repressor when it binds arabinose, would there be high or low transcription levels when arabinose is present?

Welcome to Biology.SE. Please take the [tour] and then go through the [help] pages starting with [Ask] questions effectively on this site and [edit] your question accordingly. In particular, your question isn't clear and shows no evidence of prior research. Unclear because you haven't specified what gene(s) you are interested in. Under researched because you haven't said where you've looked for answers, what you do know about the topic, and where exactly you still have questions. Please note that both of these are considered to be reasons for down-voting and closure on this site.

A repressor inhibits transcription, thus one expects low transcription when the repressor is present.
Thus, if a protein acts as a repressor when it binds arabinose, then when arabinose is present the protein will inhibit transcription.
In the typical synthetic case of AraC and pBAD, AraC+arabinose does not act like a repressor, but an activator of pBAD.  There are some awfully big caveats there, however:

If "high" and "low" transcription levels are not well-defined, you might find yourself be dealing with a different phenomenon entirely.
If you change the promoter or the genetic or environmental context, you may get different effects, up to and including changing the regulatory relationship.
Transcription is also affected by other things, and so if AraC is also affecting other things in the cell, the apparent affect might be inverted as well.

Bottom line: you need to actually define the system better in order to predict the effects.


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Is this a Box Tree Moth?
This moth has been hanging about near my house recently (South London area, October 2021) - I have seen it 2/3 times in the last fortnight. I have never seen anything this colour before which is why it caught my eye.
Its wingspan is about 30mm and this photo was taken from the inside of a window it had landed on.

Basic google searches lead me to believe it is a Box Tree Moth - am I correct?

What a beauty, do you have a picture of the back?

@CoDa Unfortunately not, it flew off before I could get one


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Do 0.2 vs 0.5 ml epitubes require different PCR machines, incubators, etc?
Eppendorf themselves and many other companies list on their website two above mentioned sizes of "PCR tubes". As I understand 0.2 ml is by far more widespread and "standard" for PCR, so do they make different machines for 0.5 ml tubes or do 0.5 ml tubes fit 0.2 ml holes? Also, what about incubators and other bio equipment that holds epitubes?

Please edit the question to limit it to a specific problem with enough detail to identify an adequate answer.

I have always done PCR and qPCR reactions in 0.2ml tubes, not aware of any blocks designed for 0.5 ml, but they may exist. But there are other form factors that you have to pay attention to. Some (but not all) qPCR cyclers are made to work with low profile PCR tubes only. Some older machines can only use high profile (standard) tubes, but adapters are available. Some manufacturers have even devised multi-instrument workflows that require both. You also have to pay attention to tube color if you're doing qPCR (when in doubt, stick with clear) and skirting on 96-well plates.

The answers are both yes and no.
Many PCR machines come with the block designed to hold both 0.2 and 0.5 ml tubes (e.g. this one from Eppendorf**), or with swappable blocks (e.g. Bio-Rad C1000**), this depends on the manufacturer, the instrument you are looking at using, and what specifications you are looking for. For some machines, such as qPCR ones, there isn't any point in having 0.5 ml tube holes as almost all the assays will be done in 96-well 0.2 ml plates or 384 well plates (0.02 ml; with a different block designed to hold these plates). A large portion of whether you can use different tubes in different PCR machines is down to how the (heated) lid closes - if the block is for 0.5 ml tubes, then the lid will not go down far enough for 0.2 ml tubes, unless it is adjustable. Conversely if it is set up for 0.2 ml tubes, the lid will go down too far for 0.5 ml tubes, deforming them and potentially damaging the machine too.
Static temperature incubators, such as thermomixers (Eppendorf brand**) often only come in one tube size (e.g. Eppendorf ThermoMixer F), but there are thermomixers that come with swappable tube holders (like the ThermoMixer C), with anything between 0.5 ml - 50 ml tube holes and even plate holders.
For tube racks, it depends again - some do, some don't. Many 1.5 ml tube racks can be flipped over to expose 0.5 ml tube holes, but it depends on the brand. You can certainly buy racks for 0.2 ml tubes (usually in 96-well format), but 10 ul - 200 ul tip boxes work well for this purpose too. 0.5 ml tubes will fit into 0.2 ml racks, but will be unstable and you won't be able to pack the tubes as densely as 0.2 ml tubes.
** I have no affiliation with Eppendorf, Bio-Rad, nor any other brand and do not endorse or imply endorsement through this post. I also do not I sell or distribute any products - I'm simply an end-user.

Here's a top-down photo of the heating block from an Eppendorf 6331 Nexus Gradient MasterCycler (source):

Most modern Eppendorf thermal cyclers have a similar block.

Are those super little ones for 0.02 ml tubes @bob1 mentions?

@beardeadclown no. count the super-small holes and you will notice that they are way less than 384. Plates with smaller volume per tube are higher density since they split the positions into 4 smaller ones when going from coarse to finer

@masterX244 I'm sorry, I didn't get any of that. What's the significance of number 384? Plates I assume mean tube holders, coarse being .5ml and finer being .2ml, and density means holes per area, but I can't see what positions do those super-small holes split into 4 smaller ones. Also, what are those 6 flat holes for?

@beardeadclown 384=4x96. Each "position" gets split into 4 smaller ones when raising plate density (except when going from a Basin plate which is a 1x1). Stepping is 6, 24, 96, 384,1536 on the standard plates which all got the same size. And i guess that the 6 flat holes are due to technical reasons.

@beardeadclown I imagine the smaller holes are to accommodate expansion and contraction of the block as it is rapidly heated / cooled. I'm not sure what the six larger holes are for, but I don't think they are meant to hold tubes

@masterX244 I think I'm starting to understand. Is there any freely available resource where I can learn more about PCR standard plates, tubes, their dimensions, design and technical specifications?

You can start by browsing brochures from Eppendorf and other manufacturers.


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Can hydroponically grown plants selectively uptake nutrients, or return excess water?
If a plant is growing with the roots in the water but the rest of the plant above water, does that plant have to absorb all of the nutrients in the ratio that they exist in the water or can it absorb what it needs and reject/return what it doesn't? My (quite basic) understanding is that plants use transpiration to create the pressure difference needed to suck up the water and all the nutrients it contains. This process doesn't allow an obvious way for the plant to either select for certain nutrients, return extra water to its roots, or reject nutrients it already has too much of. Is this because the plants actually can't, or because my understanding is flawed and too simple?
For background, I am asking because I keep aquariums. Like many people in this hobby, I grow pothos (Epipremnum aureum) in several of my tanks with the goal of removing nitrates, which can be deadly to fish in high concentrations, from the water. However if the plant is just taking in water and whatever nutrients are already dissolved into it, without any way to concentrate those nutrients or return excess water, then it stands to reason that having a pothos in the tank will not directly affect the concentration of nitrates. After all, taking a glass of 50/50 water and vinegar, and splitting it into two glasses doesn't give you one half full glass of water and one of vinegar; it just gives you two half glasses of the same mixture. There is still going to be some net effect from me refilling the tank more often with nitrate-free water to account for the water lost via transpiration, but is that it?
I have read through several questions here, but none seems to directly answer my question, or at least not in a way that I understood. While this could potentially be asked on the pets stack, I am hoping it is still on topic here. If I need to refine my question to make it fit better, please let me know.

A good search term is aquaponics .

I think most nutrients are actively transported in the roots, not dependant on transpiration rates.

I found a study from 2003, It very vaguely addresses your query, although it's not simply written: http://www.esalq.usp.br/lepse/imgs/conteudo_thumb/Mechanisms-and-Control-of-Nutrient-Uptake-in-Plants.pdf

Thanks for the pointers! @LifeInTheTrees, that does look rather information-dense. I'll do my best to work through it over the next few days and see if I can't pull out an answer


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What plant species generate the highest amount of bioelectricity?
I just want to ask which plant species produces a high amount of electricity. I'm very confused with the units that's why I don't know which species produce the highest amount of electricity. Also, can someone help me do differentiate each unit? It's just I don't have enough knowledge about it. By the way, the picture shows the species and produces power density through Plant Microbial Fuel Cell (PMFC)   Thank you...

Welcome to Biology.SE. Please take the [tour] and consult the [help] starting with [ask] for details about using this site. Your question appears to be about converting between various units, which is off-topic as it is not about biology as defined in the [help]. Have you tried searching for "power density unit conversion"? In general, we expect you to make an effort to understand the site and learn what you can on your own before posting. Please do this and then delete or [edit] your post accordingly. ——— Also don't post text as images.

I’m voting to close this question because it doesn't appear to be about biology as defined in the [help]. It is also poorly formatted (text as an image) and contains unreferenced material from an outside source.


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Could someone explain how chromosomes are counted here?

I'm not sure how chromosomes are counted in this picture. I would reason that there would be 8 chromosomes in the diploid cell, but that doesn't seem to be the case. Could someone explain?

Welcome to Biology.SE. The Biology.SE community has agreed that 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. Under researched questions may be subject to down-voting and closure. Please take the [tour] and consult the [help] starting with [ask] for details.

In particular, part of the minimum effort expected of posters is that they search on this site for similar questions and then read the associated answers. Here is one of several examples of what you can find.

Where did the diagram come from?  I'd say it is flat-out wrong, as the left shows 8 chromosomes. It shows 4 types of chromosomes or 4 pairs of chromosomes, but a total of 8 chromosomes.

Like @mgkrebbs commented, the left poster shows a total of 8 chromosomes, but only 4 pairs of chromosomes. I would dare say that you are right and that the poster is wrong. The left has 8 chromosomes, and 4 pairs of chromosomes. The right has 4 chromosomes, and they are not paired up, so the number of pairs is insignificant.

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chip sequencing
I understand the concepts of steps in Chip-seq up to DNA purification, but I don't get how one can then amplify the purified DNA samples that are once bound to the proteins.... Since sequence-specific primers are needed for qPCR, but you don't know where exactly the protein of interest binds to in the genome (that's why you do a chip-seq experiment), theoretically, how can you design primers to amplify?

Specifically, I am referring to this graphical representation of the steps in ChIP-seq
Image source: https://compgenomr.github.io/book/measuring-protein-dna-interactions-with-chip-seq.html

This question could be reduced more generally to "how does whole-genome shotgun sequencing work?" The problem of translating a high-complexity mixture of naked DNA fragments to a sequencing library is shared by many techniques. I would strongly suggest reading more on the topic of library preparation. Here is one resource.


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What would happen to the membrane potential if a cell didn't have developed relative refractory period?
If the Na+ voltage gated channels remain open instead of getting deactivated during the re-polarization period, would the membrane potential become 0 since the Na+ ions would be constantly bringing into the cell while K+ is transported out of the cell, which makes them cancel each other out in a sense? However, I feel like there is something wrong with my reasoning but I don't know which part.

Welcome to Biology.SE. You are dealing with an electrochemical potential, but only thinking about charge. Are you familiar with the Nernst potential and its extension to multiple ions? 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 also take the [tour] and consult the [help] starting with [ask] for details.

" If the Na+ voltage gated channels remain open instead of getting deactivated during the re-polarization period, would the membrane potential become 0 ? "
No, if Na+ channels remain open then the repolarization period would never happen.

Mathematically we can prove the above with the Goldman–Hodgkin–Katz voltage equation

where

approximate relative permeability values at the peak of a typical neuronal action potential are pK : pNa : pCl = 1 : 12 : 0.45 calculator

and thus we can see that the membrane potencial is much higher than the opposite for example

For a typical neuron at rest, pK : pNa : pCl = 1 : 0.05 : 0.45 calculator

and the only variable that we have changed was the  relative membrane permeability for Na+

Hypothetically speaking if [K+]i = [K+]o = [Na+]i = [Na+]o = [Cl-]i = [Cl-]o the membrane potential in that case in fact would be zero.

However that is impossible because at rest, Na⁺/K⁺ ATPase constantly moves 3 Na+ ions out and moves 2 K+ ions in, and thus these gradients are maintain, so they cannot ever be the same.

Regarding your other question
What would happen to the membrane potential if a cell didn't have developed relative refractory period?
In that case the probability of repolarization and hiperpolarization to occur is diminished and thus it's possible to assume an extreme scenario of convulsions with death associated.
Example: Strychnine is a neurotoxin which acts as an antagonist of glycine and acetylcholine receptors

on the other hand, an excessive relative refractory period scenario, would result in a difficulty to achieve the threshold and depolarization
Example: mechanism of action of local anesthetics

by applying the principle above it is possible to reversibly decrease the rate of depolarization and repolarization of excitable membranes

As a side note (or curiosity), based upon that equation we can also modify Cl- values, and in that case we can speak about the mechanism of action of hypnotics and sedatives (such as benzodiazepines)

Thank you so much of your detailed answer, I really appreciate it! If you don't mind, could you tell me why it's illogical to think that the membrane potential would become 0? I get that mathematically it changing the permeability of Na+ causes the membrane to stay depolarized but I still don't know why hypothetically speaking, in an environment where only Na+ and K+ are present inside and outside a membrane, the membrane potential wouldn't just be 0 when both voltage gated K+ and Na+ channels remain open and the total concentration of K+(inside+outside) = concentration of Na+(inside+outside).

if and only if, at rest, the Na+/K+ pump failed, then it would be possible such scenario.. but it has nothing to do with ion channels, because ion channels only open and close.. if there is no gradient then ions dont move.
Sorry in advance, I think that i got a little off topic with my answer.. I've also edited it

Thank you..I gained a better understanding now! Just to make sure, is it right to think that if both Na+ & K+ voltage gated channels just remain open, depolarization wouldn't occur, and the membrane potential would decrease (compared to ~ +45) but it wouldn't become 0, because there is always a gradient. In this hypothetical situation, Na+ would keep coming in and K+ keeps going out, but the membrane potential doesn't depend on the opening/closing of voltage-gated channels, but rather the concentration gradient of different ions. Thank you again for answering my poorly worded question


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Interactive molecular machinery visualisation software?
If I want to manually feed mRNA through a ribosome to actually see what happens step by step which software can I use to do that?
I'd like to be able to see changes in geometry and orientation at an atomic level. Maybe this doesn't exist yet, I don't know!

Welcome to SE Biology. We try to clarify questions before answering to avoid ambiguity. Your request for software for molecular visualization assumes that the necessary data input exists. This would presumably be in the form of atomic coordinates for all successive states involved in peptide bond formation of the same ribosomal machinery. Do you realize why such data are not available? If your objective is to understand the molecular interactions at each step in protein synthesis, what is wrong with original papers and standard molecular visualization of their data?

Hello thanks for the welcome. Well it sounds like I'm being too greedy and what I'm looking for doesn't exist. The atomic structure of the ribosome has been available for over 2 decades now so in answer to your question, no, I don't understand why a few successive states have not yet been recorded. The question comes from a desire to interact with and visualise molecular machinery at an atomic level, I wish to visualise the sequence of mechanical steps that happen in for example translation because visualising something is understanding something.

As you may know, one has to trap all the ribosomes in a particular state to be able to get the coords by X-ray or EM. This generally means using particular modified reagents or inhibitors. Using these techniques, we certainly have snapshots of several stages of protein synthesis — however half a dozen snapshots do not make a movie. I am sure some of the molecular viewers that take PDB coordinates allow one to align and load different structures in sequence — I imagine that you are familiar with them. If you like I can look out what different stage structures and  associated stuff is available.

I was imagining that if such a program exists it would rely heavily on modelling and simulation and not necessarily move through a sequence of perfect images. We can find so many animations nowadays and while not perfect they do a good job of giving us the general idea however I wish to go a little deeper. I have 2 jobs and I'm a student so reading through multiple papers, viewing static PDB files and trying to imagine the changes in positions and bonds all in my head seems too difficult and inefficient. I will try to create my own solution! Thank you for your help!


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How much air does a square meter of sugar beet consume per hour?
I've come across a person who tried to disprove photosynthesis (yeah, I know). The basically took numbers of record sugar beet crop in Chile (192 tonnes per hectare), and tried to calculate how much air the plant would consume in order to grow. They've ended up calculating that a square meter of sugar beet field needs 7.32 cubic meters of air per hour, and they decided that this is too much, as it is amount 14 people breath in, hence their conclusion is the photosynthesis isn't real.
Where's the error?
The numbers they ended up with were:

125 days to harvest.
That is 127 kg per hour per hectare, or 12.7 gramms per hour per square meter.
Dry mass of beet is 15% hence 1.9 grams per square meters per hour.
45% of that is carbon which is 0.86.
Carbon is 27% of mass of CO2 (12/44), since one cubic meter of air weighs 1.22 kg and that is 0.117 grams of carbon.

At this point they assumed that a square meter of sugar beet should "pump" 7.37 cubic meters of air through their leaves in order to produce this kind of harvest.
Where is the error (or more than one)?

Welcome to Biology.SE. Please take the [tour] and consult the [help] starting with [ask]. In particular, this is a site for clear, focused, well-researched biological questions — it is not a site for debunking crackpot "theories". Among other things, air is not "consumed" by plants (or animals) — you seem to be talking about carbon dioxide? Most importantly show us that you have tried to answer this yourself and how — start by providing references (to reliable sources) for your numbers.
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Oh my gosh... I would just totally file that person's belief under "useless to bother arguing with". My own father believed in spontaneous generation. I never argued with him about it. (He was not highly educated, and it would have seemed I was lording my education over him.) Pick your battles to preserve your sanity.

@tyersome, I would honestly prefer if you've answered this question instead. I've participated on se before and know the drill. Regarding the question I've checked his numbers and tried to calculate amount of carbon plant is supposed to generate myself. The thing is resulting numbers ended up close to his results, and I was unable to find an obvious error in calculations. Which means I'm lacking domain knowledge and that's why I asked it here. It is not debunking. It is looking for a missing knowledge. His conclusion is wrong, but I can't tell why. Hence I ask.

@anongoodnurse I asked because I'm unable to locate the error in their logic by myself. I don't care about battles, but I'd like to know what the error is.

Taking it slowly;
192 tonnes per hectare is
0.0192 tonnes per square metre (div 10,000)
19.2 kg per square metre over the year
0.1536 kg per day per square metre (div 125 days)
153.6 grams per day per square metre
6.4 grams per hour per square metre. (Div 24)
0.96 grams of dry matter per hour per square metre (x 0.15)
0.432 grams of carbon per ... ( x 0.45)
Derived from 1.584 grams of CO2 (div 12/44)
Using CO2 level of 410 ppm,
A cubic metre of air weighing 1.22 kg contains 0.7588 grams of CO2
The averages work out to:
the square metre of beet, per hour uses the CO2 from 2.08 cubic metres of air. (1.584 / 0.7588)
The enzyme RuBisCO captures this CO2.
The amount people breathe isn’t strictly relevant. But....
A common claim is that sedentary people exhale about 1.1 kg of CO2 per day. That’s enough to supply 28 square metres of these beets. (1100/(1.584 x 24))
This sort of maths is sometimes done for hypothetical “how much do I have to grow in my spacecraft” questions

Thank you very much. Few minor details, however:  The original poster I ran into also took day time into account (different from spacecraft, as spacecraft can have continuous light).... and I recall that sugar beet is 20% sugar and 5% pulp. Does the dry matter calculation take that into account?

I haven’t investigated the dry matter calculation; but I feel that carbon content of dry matter will be fairly constant, by a factor of two anyway. That’s a huge crop - but I regard it as referring to whole plant. Carbon content in sugar similar to carbon content in cellulose or lignin.


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A DNA extraction protocol that is equipment-less for bacterial metagenomes?
I am going into the field soon to sample a very remote environment. For reasons that are unimportant here, I would like to extract bacterial DNA from these samples in the field.
I will have access to electricity, but no heavy equipment such as centrifuge. My goal is to extract DNA from these hypersaline liquids for long-read sequencing and subsequent metagenomic analysis with a focus on DNA methylation. My plan is to filter large volumes to have sufficient biomass onto a filter from which extraction will occur (think seawater type samples).
Any suggestions for an equipment-less DNA extraction kit? Vortexing is fine.

You'd be far far better off waiting until you can get them to a lab. Given the sensitivity of the technique and the unknown abundances and likelihood of getting samples, giving them the best extraction you can is the way to go. I realize this isn't always feasible and there are reasons not to do an extraction in the lab, but it is generally better. Fixing in ethanol on a filter would be fine to preserve until you can get it to a lab.
It would depend on the scale of preparation you want to do, exactly what you want to get out of it and the abundance of the bacteria in your samples. There are a number of para-magnetic bead based DNA (AKA SPRI beads) and RNA extraction products on the market. These are highly sensitive and quite quick to perform, but you do need the magnets and beads, both of which are quite expensive.
An example of this used for environmental bacteria is the following article. They did their extraction in the lab and used a centrifuge to concentrate samples, but it could easily be applied to the field:
Byrne RL, Cocker D, Alyayyoussi G, Mphasa M, Charles M, Mandula T, Williams CT, Rigby J, Hearn J, Feasey N, Adams ER, Edwards T. A novel, magnetic bead-based extraction method for the isolation of antimicrobial resistance genes with a case study in river water in Malawi. J Appl Microbiol. 2022 Nov;133(5):3191-3200. doi: 10.1111/jam.15755. Epub 2022 Aug 23. PMID: 35946113; PMCID: PMC9804433.

The reason to do it in the field is to avoid changes in the methylation pattern changing due to temperature shock when storing. One thing I may do is flash-freeze after storing in a buffer instead, but I did hope for a baseline to compare. Is ethanol the recommended storage buffer?

@donkey ethanol more or less fixes the DNA as it was at the time of addition. I don't know off the top of my head how you would proceed to with an extraction once in ethanol, but it should be fairly easy to do. EtOH fixed samples are one of the better sources for DNA from historical museum specimens and the like.


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What is meant by 20 ul final concentration of polyacrylamide?
I am performing a phenol-chloroform extraction in which precipitation is enhanced by linear polyacrylamide. The SDS for linear polyacrylamide mentions that it should be used such that its final concentration is ~20 ug/mL.
The protocol I am following mentions 20 ul/mL. Page 21, 3rd line of the second paragraph under precipitation.
Am I misunderstanding one or both of these instructions? 20 ul/mL seems like a lot, especially given that it is sold in 5 mg/mL solution (i.e. ~5 ul/mL).
Are they referring to different concentrations, i.e. one is 20 ug LPA/mL extraction volume, and the other is 20 ul 5 mg/mL LPA solution/mL extraction?
Is there a typo in this paper?
Thank you.

Questions here should be standalone, so, ideally you shouldn't have to follow a link to understand what is going on. You should add the quote to your question - that is one massive paper and the link takes you to the webpage, which doesn't have page numbers etc, so you then have to download.

The simplest answer here is that they have made a typographical mistake in the paper and it should be 20 ug/ml.
The other possibility is that they have a stock concentration of this reagent, which they add at 20 ul per ml of precipitation volume - in which case this stock will be 1 ug/ul (1 mg/ml), assuming they use your standard concentration. When writing the paper, they forgot to convert this into standard concentration, probably (in my experience) through familiarity with it noted this way and forgetting to change it.
I note that polyacrylamide is mentioned exactly twice in the paper you linked - once in the text and once in the references, but curiously enough, not in the methods section, only in results. For completeness; the referenced paper1 adds 4 ul (20 ug) to a reaction.
Ref:

Andrea K. Bartram, Calvin Poon, and Josh D. Neufeld. Nucleic acid contamination of glycogen used in nucleic acid precipitation and assessment of linear polyacrylamide as an alternative co-precipitant. BioTechniques 2009 47:6, 1019-1022

Confirmed by the author it was a typo. ug was meant not mL.


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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.


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How sensitive are "cheap" genetic sequencers these days?
It is in the news that tomatoes with a Sl7-DR2 knockout are being trialed in England, with a view to being commercially grown and available to consumers.  While this would be legal in England, it would be illegal in Wales and Scotland, and there are concerns that the food industry may not effectively distinguish between them such that they enter the food chain in these countries.
If one was concerned about that, as I understand it, one could use the sequence given in the paper as a guide sequence (TGTTTCACTGGGCTGGTTTAGC) to detect the presence of a small amount of GM tomato in a pooled product, say tinned tomato's.  If one used a conventional next generation sequencing machine costing 6 figures of dollars this would be incredibly sensitive, such that any amount of contamination is likely to be detected (I think).  There are a number of different technologies that allow for much cheaper sequencing machines, for example the Oxford Nanopore MinION costing about $1000.
How sensitive are these cheaper sequencing machines, in relation to detecting the presence of a specific allele within a mixed population with many copies of the alternative allele?

I would hope that a similar law would be passed in Scotland, although it is interesting politically as the SNP is in hock to the Greens.

Surely you would just use PCR, not sequence the whole genome.

@PolypipeWrangler's point is correct. To answer the question not asked (What's the cheapest way to detect this sequence?), some kind of direct PCR solution would probably be the answer. You could just do vanilla PCR and run the product on a gel if you had good size standards, you could use a qPCR solution, or you could maybe test it in the field with an isothermal PCR solution.

Note: user @Polypipe Wrangler's comment about using PCR for this application is very pertinent. My answer is mostly about distinctions in sequencing technology.
Cost
An important distinction is between cheaper machines and cheaper sequencing.
You can buy time/services on instruments owned by service providers, and in that comparison Illumina remains a cheaper option (MinION is actually one of the more expensive options per sequencing run).
The upfront cost of the instrument, while relevant, is ultimately not the practical point of comparison between these different methods.
Sensitivity
Either instrument is roughly comparable at this point for finding a given sequence per sequenced base. Illumina is still more accurate, but with bioinformatic advances you can overcome that limitation pretty easily.
The advantage of Illumina comes in that it provides many more sequenced bases- that is where your sensitivity comes from. You will have to run a lot of MinION flow cells to get comparable numbers of reliable bases compared to Illumina.
For specific information about estimating limits of detection for high-throughput sequencing, see here. The short version is that detection of a given chromosomally integrated sequence in a sequencing run should be excellent, assuming that the sequence's copy number is comparable to chromosomal copy number.

I would start by pointing out that PCR-based detection methods are generally found to be more sensitive and specific than sequencing for applications like this one. (see reference). While qPCR methods are still considered the gold-standard for genetic detection, digital PCR (dPCR) has several advantages and can often achieve greater sensitivity and specificity.
Ultimately, the sensitivity depends on the sequencing depth and assay design. Speaking from experience in methods development and validation for pathogen detection in complex biological specimens, even massively high-throughput next-gen sequencing systems like Illumina MiSeq or HiSeq platforms don't really match the sensitivity of PCR-based detection method unless you implement them with some form of target enrichment, like hybrid-capture probes.
And in terms of cost, a standard endpoint PCR or qPCR will probably win out each time, if you're only interested in the one target. Especially if you factor in the target enrichment, which usually involves some kind of amplification using a thermocycler like the ones you use for the PCR detection anyway. The one area where a targeted sequencing approach might make sense is if you were screening for large panel of different targets in each sample, which doesn't seem to be the case here.


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Why do so many pollutants affect the estrogen receptor?
We make loads of chemicals, and release them into the environment for reasons.  We have loads of hormone receptors.
There are many instances of human made chemicals affecting the estrogen receptor, currently in particular hair straightening chemicals associated with higher uterine cancer risk, and also in other areas of the internet that I am loath to link to.  I can think of no other hormone or neurotransmitter receptor in the human body that is affected by so many human made chemicals, at least if one follows the media.
Why is the estrogen receptor affected  by so many human made chemicals, and other hormone receptors apparently not?

Probably, many other endocrine receptors also get functionally disrupted by small molecules people manufacture. I think there's a dearth of research studies designed to find them. So, many molecules disrupt the estrogen receptor because researchers are biased by the Streetlight Effect. The absence of evidence for disruption of other endocrine receptor functions is likely insignificant.
Sexual development is probably an endocrinological "canary in the coal mine" rather than being more sensitive to exogenous small molecules than other endocrine contexts. The estrogen receptor gets its pathophysiology described so extensively because of the way teratogens are characterized; medical scientists are experienced with studying the developing fetuses of lab animals, and perturbed sexual development is measurable with widely known correlates (like anogenital distance) that work as popular response variables.
Once a hypothesis is generated, say that molecule A disrupts development by binding to a sex hormone receptor, then followup experiments lead to experiments where binding affinities get measured. So by directly incubating molecule A and a receptor, a binding affinity gets measured. But this is rarely done in the presence of competing ligands or receptors.
Looking for references to explain this, it's difficult because it's a kind of subtle background truth about experimental designs. Here's a quote from a paper that quotes a bunch of other papers (albeit in a different context of protein:protein binding):

[H]igh affinity does not guarantee high specificity, which is controlled by either partner binding to other proteins discriminatively (Bolon et al., 2005; Grigoryan et al., 2009; Havranek et al., 2003; Janin, 1995; Kortemme et al., 2004; Shifman and Mayo, 2003). Paper on specificity in protein:protein interactions

So, for example, all the 'estrogen analogs' that are measured to bind to an estrogen receptor also have affinities for an androgen receptor, an opiod receptor and a glass surface, but it's simply not the usual research practice to characterize a wide range of affinities. Imagine the technical requirements of an experiment of, say, studying the competitive binding of a mixture of estrogen, cholesterol, and porphyrin binding estrogen receptor, let alone a competitive mixture of receptors.
If a ligand does bind well to a receptor, people tend to assume it does so specifically and (patho)functionally but this is oversimplified.


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How dependant is food calorie content on cooking method?
Most food available in the supermarket is labelled with nutritional information including Energy expressed in kcal/kJ and is frequently given to 3 or even 4 significant figures.  This is defined (by the FDA as "Calories provide a measure of how much energy you get from a serving of this food".
There is a theory popularised by the 2009 book Catching Fire: How Cooking Made Us Human.  I have not read the book, but my understanding is that it postulates that the increased nutritional value of cooked food allowed us to evolve multiple features that made us the dominant species.  This would seem to indicate that the calorie content of food is highly dependant on the preparation method, in particular the presence and extent of heating of the food.
How accurate is nutritional information in the presence of different cooking methods?  How similar is the calorie content between rare and well done steak, or coleslaw and Irish cabbage?


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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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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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Which test can be used to diagnose typhoid fever
The only thing I found was the widal test, which is no longer an acceptable clinical method. I can't find much information about this anywhere else other than the test needs to analyze samples of blood, poo, and pee. Is there a specific name for this test?

Welcome to Biology.SE! Please take the [tour] and then go through the help pages starting with [Ask] questions effectively on this site and [edit] your question accordingly. In particular, it is good to check multiple sources before posting — have you read the wikipedia article on typhoid? Thanks!

I have voted this question as off-topic for SE Biology as it is merely asking about current clinical practice rather than about the biology underlying a medical condition or test. It might be on-topic on SE Medical Sciences.


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Why does wheat consume more water than rice that is, unlike wheat, flooded?
Why does wheat consume more water than rice that is, unlike wheat, flooded? I thought rice was the villain of the piece when it comes to water consumption.

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Is there any evidence ethylene (gas) has any detrimental impact on human health?
Is there any evidence ethylene (gas) has any detrimental impact on human health? The only thing I was able to find is that the IARC doesn't consider it a carcinogen. But carcinogenesis is not the only way a substance can afflict human health, is it? The ATSDR only has ethylene glycol and ethylene oxide. But maybe it's oxidized after being emitted so the latter is relevant, isn't it?


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How do I calculate a "full-life" from a half-life of substance elimination?

The half-time of carbon monoxide disappearance from blood under normal recovery conditions while breathing air showed considerable between-individual variance. For carboxyhaemoglobin concentrations of 2–10%, the half-time ranged from 3 to 5 h (Landaw, 1973); others reported the range to be 2–6.5 h for slightly higher initial concentrations of carboxyhaemoglobin (Peterson & Stewart, 1970).

(from here)
To get a "full-life" (or the best approximation of it), should I multiple it by 2 or 6.64 (99%)? Is carboxyhaemoglobin's "full-life" in bloodstream 4-13 or 13-86 hours? That's a huge difference

There is no such thing as a "full-life", because in most relevant mathematical models (the simplest and most common being first-order exponential decay), the time required for all the the substance to be gone is infinite.
For example, the half-life is marked on the following graph as $t_{1/2}$. Is two times the half life a "full-life"? What must $t$ be for the quantity (concentration, number of particles, whatever) to become zero? Based on my first paragraph you should be able to answer these questions.

What you can do, given data or an equation, is calculate the amount of time required for the quantity to drop to a specified amount. For instance, you could decide that you'd like to know the time it takes for the CO concentration to drop by 90% or 99% or 99.9% from the initial level. A rule of thumb (which can be derived from the exponential decay equation) is that the quantity will be depleted by around 99% in approximately seven half-lives. You could derive a similar relations for any desired percent. Which percent to choose for the cutoff of interest is at the discretion of the person doing the analysis, and different fields and contexts have different standards. The substance will never be "completely" gone, but there may be a meaningful threshold beyond which it becomes negligible.