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Can a blood type O be born from AB and A parents?
I have a basic understanding of genetics, and I'm really puzzled by this. My grandma's blood type is A (I don't know if it's heterozygous or homozygous) and my grandpa is AB. Yet my mother is type O.
Grandma says that my mom is 100% my grandpa's child, so how did it happen?
I have a few hypotheses regarding this. Either my grandma is lying, which I find to be really implausible, or there's some genetical anomalies either in my mom or my grandparents.
I suspect that grandma is heterozygous, having the IAi genes for blood type, and grandpa has a weird case of IAIBi alleles, coming either from a failed disjunction which caused both alleles for A and B to be present in one sperm cell, and the i allele to be present in the egg, resulting in the IAIBi genotype. Then, when grandpa had my mom, both i alleles got combined and out came my mom with O type.
My other hypothesis is that mom is heterozygous, either IAi or IBi, but the dominant allele underwent a deletion (or something else) and was inactivated, and as such only the recessive allele remained, giving her type O blood.
Could I get some insight on how this happened? I can't seem to find anything related to it on the internet.

Never discount the prospect that someone in the lab screwed up.

Maybe have a look at the Bombay blood group. It is a pseudo 0 blood group.  https://en.wikipedia.org/wiki/Hh_blood_group

The simplest probability is that is wasn't grandpa's sperm that did the job and that grandma either "jumped the fence" or was raped, and either way was afraid or ashamed to admit the truth.

One thing for you; the "failed disjunction" hypothesis is testable; if true a genetic test would report your mother to be the child of her father's non-extant brother.

As @KaPy3141 mentioned, other genes such as the Bombay blood type may suppress the A and B alleles (epistasis). There was once an episode of a soap opera that had this exact plot.

It can't have been an error in the lab, because both my grandparents have been blood donors for years. If they messed up the blood types a lot of people would've died. Thing is my grandma and grandpa aren't on good terms anymore, nor am I with my grandpa, so I can't really have them get genetics testing. It's not that important either way, I'm not trying to test my grandma's fidelity, I was just curious if such a phenotype would be possible from this genotype.

Did any of the involved persons have a bone-marrow transplant or suffer from auto-immune diseases? Then maybe we should also consider the possibility of a change of blood-type.

The ABO blood type is controlled by a single gene (the ABO gene) with three types of alleles inferred from classical genetics: i, IA, and IB. The IA allele gives type A, IB gives type B, and i gives type O. As both IA and IB are dominant over i, only ii people have type O blood. [1]
So from this standpoint, normal Mendelian genetics cannot explain the blood-group 0 offspring, given the AB parent.
However, besides errors in testing, the (pseudo?) 0-bloodgroup could be explained by the Bombay phenotype. I don't know the incidence rate in your region, so I can't comment on the likelihood.
To give theoretical background, this would be considered a pseudo 0-phenotype, as the individual genetically does not need to be ii. Instead, the individual lacks another effective enzyme that is required for the A or B alleles to be effective; the blood-type specific sugars (N-Acetylgalactosamin|Galactose) cannot be attached to the red blood-cell membrane, as the anchor of these sugars (called H-substance) is not present.
Finally, to address your hypothesis of a gene-duplication followed by a gene knock-out: There is no need for a preceding duplication. The more probable cause is a single point mutation (or multiple) near the active center of the inherited A/B enzyme, or a nucleotide insertion/deletion in its coding region, causing a frame-shift. A completely (not partially!) inactivated A/B enzyme would effectively represent a novel 0-genotype.

Your link says its incidence is typically 4 per million, up to 1 per 10,000 in select areas/populations. IMO, unless there's a family history of it, it still seems likely that somebody's lying

Novel gene knockout is also possible; I have no data no likelihood.

incidence reports are based on tested cases - so as testing rates increase the reported rate will rise to the true rate.  Microchimerism is another possible cause.

@Joshua. I agree, I edited the text!

@Punintended that doesn't follow at all.  4 per million means that on average 4 people in a randomly selected sample of 1 million will have the genotype. Since we live in a world of several billion people, it is expected that some will have this genotype. Therefore, the fact that it is rare doesn't tell us anything about whether the grandmother is lying. Some people do have this, so why not the OP's mother?

@terdon Of course some people do have this, and if this disorder is somehow confirmed, the probability is one.  But without prior knowledge, based on the average incidence in the world (or even the subpopulation), the odds of her having it are indeed between 1/250k and 1/10k.  That's kind of the point of "average incidence"

@Punintended yes, but the odds alone are meaningless without taking into account the size of the sample pool. Essentially, here we are asking "what are the chances that a random person selected out of a pool of ~8 billion is one of the 4 per million" and that is a very different question to "how likely is it that this person is one of the 4 in a million".

@terdon If you have 100k people and an incidence of 4/million, stochasticity kicks in and the odds change. That's not the case for large sample sizes, like an n of 8 billion. 4/million * 8 billion is 32k. The odds this person is one of those 32k (given no other knowledge) is 32k/8b, or the incidence rate

@Punintended but we have other knowledge. So that’s irrelevant.

both my grandma and my grandpa have been blood donors for years, so we can safely rule out mistakes in testing. Thanks for your answers!

@PateArdealul and your mother?

@Tim she did donate blood a few times too. Most of my family got to know their blood type by donating blood.

Two highly unlikely, yet possible ways this could happen:
One extremely unlikely possibility is that grandmother was born O.  Grandmother's current A typing could be the result of interplacental stem cell transfer from bearing an A child and the mother is an XX clone of the grandmother. How similar in appearance are grandmother and mother?
A less rare yet still doubly-unlikely scenario is that the grandmother and grandfather are both chimeras and at least some of the grandmother's eggs and at least some of the grandfather's sperm are O.  We are learning that human chimeras are more common than we used to think. https://en.wikipedia.org/wiki/Chimera_%28genetics%29

Given that your grandmother is type A and your grandfather is AB, there is very small, minute chances that your mother is in fact the biological child of one or both of them.  For your grandmother to say that she is 100% your grandfather's child, maybe he had her with someone other than your grandmother, and due to unfortunate events, that woman died giving birth or shortly thereafter.  It may hurt your grandmother to admit that. Or, she could have the same views as many others, like myself, and recognizes that there is a BIG difference between being a father and raising a child, and just being a sperm donor and disappearing. There are many different situations that could be possible, that your grandparents may not be ready to admit for whatever reason, or it could very well be a simple mistake at the lab who did the testing. But does it honestly matter if she is the biological child of both grandparents, or whatever the truth may be? Will knowing the answer change anything?

Welcome to Biology.SE! On this site, answers are expected to be fact based (and are much more likely to receive a favorable response they include supporting references (primary literature is best)). Your answer is entirely opinion and thus is not appropriate here. ——— Please retake the [tour] and then consult the [help] pages for additional advice on [Answer] effectively on this site and then delete or [edit] your answer accordingly. Thanks!

Your theory of "biological grandmother is not the grandmother" doesn't really solve anything; the grandfather is AB, so the child would be A, B or AB, even if the biological grandmother was O. You'd still need to explain away the grandfather's lack of O managing to contribute an O, making the possibility of an unknown biological grandmother non-explanatory.

Did not realize that I needed to specifically state that there is a possibility of the mother being adopted and never told, thus explaining why she has neither of their blood types. My apologies, I will be sure to be more specific next time.

Please [edit] your post to make it better or, if you don't feel that is worth your time. then please delete it (click on the "delete" next to "edit" under your post. Thanks!

authors: Bob Jarvis - Слава Україні


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How can I change the payload of a bacteriophage used to transform E. coli?
I was looking at bacteriophages and how they're used to transform E.coli. While the whole process of how a bacteriophage works makes sense theoretically, I wanted to know how one goes about changing the default payload of a bacteriophage with the payload of interest (engineered plasmid) in practice?
Clarification:
payload - Genetic material that the phage inserts into the bacteria
default payload - Whatever genetic material the phage inserts into the bacteria naturally
payload of interest - Engineered plasmid that I want to insert into the bacteria

I would recommend looking at some methods sections. Takara has a lengthy guide for using different kinds of viral transduction methods in mammalian cells, which is a little different but quite similar overall, that may have some helpful low-level information. Here is a review that may be helpful, focusing on engineering methods for bacteriophage, with particular relevance to infectious disease applications. They go ove

Can I ask why you want to use bacteriophage as opposed to electroporation or heat shock? I can't say I've ever heard of someone using phage to deliver DNA. I know I said it in my comment on your last post about electroporation, but it was more of a mental spitballing as to why lentivirus wouldn't work on E. Coli.

@yp66t89 I am actually familiar with some projects that were using phages to do rapid diagnostics type assays. I only made the connection once you mentioned retroviruses. But that got me curious about how you actually setup the phage payload in the first place.

I am currently using a 'phage integrase' system to insert my gene of interest into P. putida and it is definitely a seamless process with high efficiency so I highly recommend it!
As for your question, the method through which we integrate the gene of interest using the phage integrase system is simpler than you think. We do not change the payload of a bacteriophage. We simply electroporate with two plasmids, one that expresses the BxB1 integrase and another plasmid that has our gene of interest that we want to integrate into the genome of P. putida. This integration is highly efficient, and it operates through a serine recombinase that recognizes specific attP and attB sequences in the plasmid that contains my gene of interest and native genome of my engineered strain. This method is significantly more efficient for engineering certain strains (typically non-model organisms) for genome integration purposes.
References for additional information on the system for your review:
Here is a paper that describes how we use the phage integrase system to effectively edit the genome.
https://www.sciencedirect.com/science/article/pii/S2214030116300438
Here is the paper that discovered and characterized the mechanism this system operates through.
https://www.sciencedirect.com/science/article/pii/S1046202310003142?via%3Dihub

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How does the Endoplasmic Reticulum scale with Cell Volume in Epithelial Cells?
I am working on a mathematical model of a biological tissue (drosophila pupal notum; an epithelial tissue) where the tissue is built up from cells all described by the same cellular-model. The tissue is generated using a somewhat random process, and so each cell in the tissue has a different size (volume, surface area, etc.)
The model focuses on the movement of particles both between cells as well as between the cytosol of each cell and internal stores (like the Endoplasmic Reticulum (ER)). In order to use the same parameters across the entire tissue, I need to know the ratios of various volumes and surface areas, such as the ratio of the volume of the cell to the volume of the ER, or the ratio of the volume of the cell to the surface area of the ER.
So my question is, if we have a cell of some volume $V_1$, and we look at another cell with volume $V_2$, what can we say about how the volume of the ER of cell $1$, $V_{\text{ER},1}$, or the surface area of the ER of cell $1$, $A_{\text{ER},1}$, compares to the volume and surface area of the ER of cell $2$, $V_{\text{ER},2}$, and $A_{\text{ER},2}$? For example, would we expect $V/V_\text{ER}$ to remain constant across all cells in the tissue? Or maybe we would expect $V/A_\text{ER}$ to be a constant across cells in the tissue? Or maybe both of these ratios follow some other sort of scaling law?

Do we speak about volume-ratios within the same cell-types at different cell-states or do we compare different cell-types? I think this question is too specialized and you won't receive an answer. It could help if you provided allowed generalizations; i.e. you tell us what cell-types (or what one specific cell-type) you are interested in, and I am quite sure there will be people who can tell how those ratios behave in similar cell-types/tissues, that are not exactly drosophila pupal notum.

@KaPy3141 I'm not thinking about various cell-states. As for cell type, do you mean like epithelial cells?

But what would case different cell-volumes? Different points in the cell-cycles? Different cell-types that naturally have different sizes?

@KaPy3141 Or just cell-cell variability. The tissue I'm considering has the same cell type throughout.

Ok, I see. At least that is now defined. Maybe someone has data.

Please consider this "answer" as an enlarged comment. I can't answer your question, but it's possible that my speculations or considerations could be helpful in your particular situation, in the early phase of building a model.
First of all, I doubt that general formulas that link cell-volume to ER-volume of different cell-types can exist. Dependent on the specific cell-type, (dependent of the protein production need in the rough ER), the ER/total ratio can strongly vary (i.e. compare neurons (strongly protruding ER), red-blood cells (no ER), pancreatic cells (large ER), endothelial-cells..). Thus, any correlation may just be chance, without the general meaning that you truly seek.
Therefore, you should focus on such volume-correlations within the same cell-type at different cell-cycle stages.
In this context, I can think of mathematical considerations, that could help your model: When cells divide, they effectively half their membranes (membrane-lipid count). Mathematically, this means that the overall volume of the cell should reduce to around 35%. In contrast, the volume of the ER as a highly condensed structure could scale linear with its surface area, if we see it as a long tube that gets separated. From this, you could build an easy mathematical model, in which ER volumes are relatively bigger, when cells are small.
However, according to cell-size measurements this is not true; cell-volumes get halved [1] when cells divide, which should indicate relative stretching of the membrane. Unfortunately, I could not find any data on how the stretching of the cell-membrane relieves over time. Further, the volume (quite) linearly increases over time until the next division can take place [1].
This means the cell-membrane keeps a constant weight/V before and after cytokinesis, while the surface-area increases by root 2. Given, that the ER-membrane-mass is also halved when the cell divides, and there is no apparent mechanism that should cause swelling, it is still quite difficult to predict how the ER volume would change. Again, the ER is no simple sphere, but has a highly collapsed shape.
However, I found this source, that deals with changes in shape and volume of the ER during the cell-cycle. Apparently, there is a membrane exchange between nucleus and ER and the ratios in volume between ER and nucleus is constant throughout the cell-cycle. You can study this yourself in more detail, but I would take this observation as an indication, that the ER volume behaves as would be inferred from a mathematical standpoint; to the third power of the diameter of the cell.
To sum it up, I initially expected smaller cells seem to have relatively larger ER volumes, and I suspect, that you had a chance to fall for the same trap. However, if the volume-ratios are not constant, they should rather behave the opposite way: ER takes less cell-volume after division, when the cells are small.

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Saturated Mutagenesis Screening
I am hoping to mutate the active site of the enzyme I am researching that has 5 residues in proximity with the substrate. I am wondering how many colonies I'll have to assess to theoretically sample all the possible mutant combinations?
I am guessing that just screening the number of possible combinations will not be very successful because I am likely to sample a particular mutant more than once?
I have been looking at 'NNK Saturated Mutagenesis' which uses the redundancy in the genetic code to be able to create all residue encodings where N = A/G/C/T and K = G/T, while limiting the number of repeated residues from synonymous codons.
Are there any other ways of performing this sort of mutagenesis with better codon compression?

It depends a bit how you are generating the mutations and the residue combinations and what your goals are. e.g. do you expect the genotypes for each mutant position to be independent? Do you want to sample every amino acid at every position, and then do combinatorics of the $20^5$ combinations across the 5 positions? That's a lot of combinations to sample (3.2 million)! Why not a greedy approach, wherein you saturate mutations at each residue independently, and then construct combinations based on whether substitutions have an effect/seem interesting?

@David 'Synthetic biology is a field of science that involves redesigning organisms for useful purposes by engineering them to have new abilities' - genome.gov,  I am looking to re-engineer my enzyme for new purposes

An enzyme is not an organism.

@MaximilianPress It is a lot of combinations but I feel it is neccessary given the dependency of residues within the small space of an active site. I am not sure you would get the theoretical global fitness optima with a greedy approach but I think this is an interesting idea for maybe tackling even larger libraries

@MaximilianPress Greedy definitely isn't effective for some of the related problems like RBS and promoter engineering, and I won't expect it would be effective for active site engineering.  One might reduce the space by considering AA permutations rather than NA, but I'm not sure if there's a good mutagenesis protocol for that.

@David An enzyme is not an organism, but if you want to give a new chemistry-related function to an organism, a typical starting point is to select enzymes to add to the organism to implement the desired function. Often, because you're transferring the enzyme from its native context to a new context, you need to engineer it, e.g., with techniques such as the one being asked about.

@jakebeal Tags are meant to indicate what the question is actually about, not what it might lead to. They are meant to help people find questions in their sphere of interest. On the basis of your argument you could add "cancer research", "covid", "evolution" and anything you thought an engineered protein might be useful for. But it would not be useful to people interested in these fields. It is not always easy to find the best tag — there is no tag for mutagenesis here, so in the list of relevant tags I have put in my edit I have put mutations. It doesn't need to be perfect, just not wrong.

@David I appreciate the careful explanation of your reasoning, which is much more constructive than your now-deleted comment that led me to my prior interpretation of your reasoning. I particularly think the "protein-engineering" tag is useful. In my own thinking about taxonomy, I would tend to consider that a subset of the "synthetic-biology" tag, but can see how others with different backgrounds could evaluate their relationship differently.

This sort of method is indeed quite useful and frequently used in synthetic biology: I've used a similar approach before to generate 5' insulators for promoters.
Calculating the exact theoretical likelihood of what you want, so far as I understand, is a complex combinatoric equation that I wasn't able to get a nice closed form on with a few minutes of work. For your purposes, however, I expect that a reasonable estimate will do as well, and that can be made much more easily by modeling the population of each combination as an independent series of biased coin flips, i.e., a binomial distribution.
Consider a situation with $n$ potential combinations that you attempt to cover with $k$ random mutants. If we focus on a particular combination, we can see that each random mutant has a $\frac{1}{n}$ chance of hitting that combination. The whole set of mutants is thus a series of $k$ biased coin flips, with a probability $p_0 = (1-\frac{1}{n})^k$ of getting no mutants that hit that combination. This means that the probability $p_c$ that the combination is covered is $p_c = 1-p_0$, expanding to: $$p_c = 1-(1-\frac{1}{n})^k$$
With a large number of potential combinations and a large number of mutants, we can make a reasonable approximate by treating each combination as independent. In this case, the probability that all combinations are covered is the product of the probabilities that each combination is covered independently. They aren't actually independent: each combination that gets covered slightly detracts from the ability of other combinations to be covered. An estimate using an independence assumption is thus an overestimate of the actual likelihood, and the higher that $k$ is the closer the estimate should be to the exact probability. In short:
$$p_{all} < (1-(1-\frac{1}{n})^k)^n$$
This smells like the sort of equation there ought to be a nice identity for, but I don't know the identity, so I'll just give a couple of examples of calculating with it:

NNK = 32 combinations, 50 mutants (n=32, k=50): $p_{all}$ < 0.07%
NNK = 32 combinations, 100 mutants (n=32, k=100): $p_{all}$ < 25.5%
NNK = 32 combinations, 200 mutants (n=32, k=200): $p_{all}$ < 94.6%

As you can see there's a sharp transition as the coverage count increases.
Important caveat: the theoretical value may not give you what you want. In the paper I linked above, we used two degenerate primers with 18 N bases to inject 36bp of random material, which means we had $4.7x10^{21}$ possible combinations and should never have seen a repeat --- except that sequencing showed that we did indeed get several repeats out of only a few hundred colonies. Thus, biases in the biology may lead your statistics to be skewed from what the theory suggests.

CyberDope was first written at MIT, then again at Kairos, then again by me (privately) in Mathematica.  The program gives the best "dopes" for a target set of amino acids.  You will need someone who has Mathematica to view this program in ".nb" format and to run it.  I would further suggest they upgrade the program into Mathematica version 12.  I just now uploaded a copy to the Wolfram cloud (public):
CyberDope
The original publcation is:
Arkin and Youvan
Twenty years ago, the work was at this stage:
https://www.researchgate.net/publication/228715857_Directed_evolution_and_solid_phase_enzyme_screening
If you just want to "eye-ball" a good dope, look at this figure, circle the amino acid residues you want, and then look at the three nucleotide axes:

(That's from Wikipedia's Genetic Code article)

@David - Please have another look.

@jakebeal - I was not on this forum when you posted.

@Youyan I'm having a hard time understanding how this answers the original question. It looks like relevant software, but I don't see how one gets from this to a number of random mutants.

@jakebeal I saw you had mentioned NNK mutagenesis, which will increase in complexity with 32^n, where ^ is exponent and n is the number of amino acids you are simultaneously changing in the protein.  For a given n, CyberDope can help you reduce the base of 32 to a lower number.  That effects the number of mutants which you will have to screen if you select a good target set of amino acids.  Lets say you want to try the aromatic residues at a site. Looking at the cube (no program), U(UAG)(UG) gives you F,Y,W with a nucleotide complexity of 6.  NU(GT) is hydrophobic, NA(GT) is hydrophilic.

@JEJS Sorry I got OP confused with users who had answered.

Thank you for the interesting reference. As a matter of process, however, if you want to respond to an answer, the recommended way to do so on StackExchange is via a comment on that answer, rather than another answer that may not be addressing the question.

The OP is dealing with combinatorial complexity, too.  So they might want to look at this figure if they are mutagenizing at the second position of the codon.  Again, from Wikipedia (Genetic Code)


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Does parental conflict lead to genes combining important functions with functions only advantageous for one of the parents?
In a sitation of a mother-father conflict of interests, the mother might use epigenetics to turn off some genes only advantageous for the father's genes and not her own. I thought a logical father's contra-strategy would be to bind the functions only advantageous to him to some of the genes crucial for them both, so that the mother couldn't turn these genes off.
Is there any evidence for a similar mechanism of "taking some genes hostage" in a situation of a conflict of interests?

While epigenetics is something the mother organism may control to some extent, I don't see how the father could do anything of the sort. Binding genes together requires genetic changes that take many generations, i.e., "father" in this case is not an organism but indication of the way the gene was transmitted.

@Vadim I broadened my question to any parent because I imagined it could be the case if the father's genes affected the chances of the fetus being miscarried.

It sounds like this question is asking about imprinting. However, you don't specifically mention that term so perhaps you are unaware of the research on this subject? If so, there was a relevant review published last year in PLOS Genetics. A "Learn Genetics" page and  "journal club" article in Nature could also be of interest.


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Can someone identify this jellyfish?
Recently, I saw this video but I couldn't find out if it's CGI or if it's an actual jellyfish. If it's real, what explains its beautiful bioluminescence pattern? And could the output from the ROV thrusters rip it apart (in the video description it says so)?
Photo:

Location: off the East coast of Africa.
Size: unknown, but it can be compared to the objects in the surroundings.
Additional info: footage taken in 2013.

The video you linked to, and from which you got the still shot. is real.  There are  hundreds of such videos now that Underwater Remotely Operated Vehicles (ROV) are more commonplace.

...could the output from the ROV thrusters rip it apart(?)

A working class ROV needed for deep ocean floor exploration can be the size of an SUV. There's no reason to believe its thrusters can't generate the shear forces needed to (sadly) tear a delicate jelly apart.

...I couldn't find out if it's CGI or if it's an actual jellyfish.

Although not a jellyfish, it is similar in being delicate; it's a ctenophore or comb jelly. The refraction/bioluminescence pattern is common in ctenophores.

And what dazzling colors these creatures display! The cilia of the ctenophores refract light in iridescent waves. Certain species are even capable of producing light, like fireflies of the sea!

For more information and videos, this short Youtube video might help. Or for a written explanation, this article, Introduction to Ctenophora  (https://ucmp.berkeley.edu/cnidaria/ctenophora.html) is good.


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What is the mechanism by which seeds stay alive while in storage?
I know folks who store seeds for months and sometimes a year before planting them. What is the mechanism by which they stay alive?
Also some associated questions:
How do the cells in the seed stay alive without resources for so long?
How long can the seeds stay alive in storage?
Does storing them long affect their growth?

What resources have you checked on seed dormancy? You’ll likely have to narrow the scope of your question for it to receive a good answer.

I suggest a little preliminary research.  But a year is really nothing for seeds: there's a date palm in Israel that was grown from a 2000 year old seed.  With a little assistance, plants were grown from 31,000 year old Silene stenophylla seeds.  It's also perfectly normal for seeds in the environment to grow after many years, wen conditions are right, e.g. a rainstorm for desert ephemerals, a fire for seeds that require smoke exposure to germinate, or just being exposed to light when I till my garden :-(

Seeds are made from cells in an amorphous metastable superviscous state because the cell's cytoplasm becomes a solid matrix of hydrogenated oil and sugar. The fats also contain a lot of anti-oxidants, so that ambient oxygen that can affect the dormant cells is absorbed by buffer chemicals.
The sugars and oils in the cell cytoplasm (the inside of the cell) become hydrogenated, and change into margarine and then become solidified fat and solidified sugar. Then the consistency of the cell at that stage becomes glassy because it's so solid, which is what you can see if you try to crack a peanut or a lentil, it has a glassy surface. The cell fats can rapidly become liquid again if they absorb water and change their chemistry.
references and images
It's not very different from microbe dormancy, so it's like an ancient biological trick that cells have been doing for billions of years.
The cotyledon is the future baby leaves of the seedling, the radicle is the root and the plumule is the leaf shoots, they are all made of cells that are in a solidified dehydrated form made of fats and sugars and special chemicals that resist ambient water and oxygenation in a controlled way.


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Why do nearby cells have similar meiosis periods?
A few days ago, I conducted an experiment to observe the meiosis of rye ears. During the experiment, I observed that cells in similar locations have similar meiosis period. Can anybody tell me why does this happen? Does the checkpoint of a single cell affects the checkpoint of nearby cells?


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Is human zygote unicellular or multicellular
Is the human zygote which is formed by fusion of sperm and egg a unicellular structure? Bcz at some places the 2 celled stage which is formed due to cleavage has been labelled as the zygote. So is the zygote 2 celled stage or the unicellular structure which is formed initially.

first three lines of this page. https://www.google.com/search?q=zygote&oq=zygote&aqs=chrome..69i57j69i59j69i60&sourceid=chrome&ie=UTF-8 The zygote's single cell ... closing question

A zygote is produced when the gametes, a sperm and egg cell, combine with each other. This is unicellular. That one cell then duplicates into two cells, and then four cells. People don't really have a need to name this stage, but anyone would understand it if you just called it two-cell or four-cell within the context of zygotic development. As more cells are created (16+), the solid mass of undifferentiated cells is called a morula. This occurs around 3-5 days after fertilization. This then becomes a sort of hollow sphere of cells. These cells are now called blastomeres and the whole structure is called a blastula. When you hear a cell name with "blast" in it, it typically means it is a precursor to some other type of cell. In this case, those 16 or so cells are the precursor to a future human's entire body.


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How pricey does the media need to be for a typical bacterial culture experiment?
I'm considering various lab supplies for a protocol we're going to be running, and have been struck by the remarkable difference in price for different quality levels of the same basic substance.
Sigma will sell you water, for example, for anything from \$15/L to \$500/L and even over \$10,000/L, depending on what level of purity is guaranteed. Obviously, one doesn't want to use inferior tools and guaranteed unreliable results, but also it would be foolish to use a reagent that's far pricer than one's actual tolerances.
Most of the actual protocols that I encounter, however, don't seem to actually specify what grade of purity they need, and I'm trying to figure out how to navigate the problem. Trial and error is definitely not a good option, since (a) it can waste a lot of time, and (b) good and bad results in a trial don't actually tell you what you need (a good result could be an unusually good batch of poor reagent; a bad result could be from an entirely different problem). Asking the authors is also not a great solution, for a number of similar reasons.
To make this concrete, let's consider a very simple and typical case that I am currently encountering. The protocol I'm setting up for right now cultures E. coli in LB media. Sigma will sell me LB media powder as "quality 100" L3152 for \$139/kg and "quality 200" L3522 for \$182/kg. Both say they are "good for E. coli".
Is there any reason not to pick the cheaper media? How should one decide how much quality their media needs to have for a typical bacterial culture experiment?

It looks like Merck have different levels of quality control for their products. MQ 100 being the lowest up to MQ 600 being the highest. From what I can see the MQ200 product here has defined composition (of salt, tryptone and yeast extract in g/L) whereas the cheaper MQ100 product does not appear to have its composition ensured.
I am sure your cultures will grow in cheaper media but perhaps you would theoretically lose out on some replicability and perhaps if you were doing metabolically focussed experiments you would want to be certain of your media composition.
As for water, I have always used distilled water which is then autoclaved after you add the media with no problems.

What quality level do you tend to use in your own experiments?

@jakebeal I am fairly sure the composition is ensured - I think it would perhaps be best practise. But again, definitely not required for microbial growth or is really likely to have a meaningful effect on your results - I guess scientists can be quite pernickety about these things though!

The answer depends on the application and level of oversight/compliance you require. Most research labs I've seen use de-ionized or even reverse-osmosis water (from a tap) for most culture media applications. But when I was doing trace-metal analysis, I had to use higher grade water for almost all steps in my experiment. Mostly, we needed a certification with each batch that our specific analytes were below acceptable levels (usually around 1 part per billion). But even "trace-metal" grade reagents often contained iron in the part per million levels, so we had to order even higher grade (pricier) water when we wanted to look at trace-level changes in iron metabolism.
One challenge is that there are few industry-wide quality grading systems. Most of them are specific to a given manufacturer, so it takes a bit of homework to understand what the mean, specifically. For example, it seems like the M-Clarity system used to define "quality" in the link you shared is a Sigma-Millipore specific program designed to help customers with product selection for regulatory compliance. The MQ 100 - MQ 400 levels all appear to follow the same industry manufacturing standards (ISO 9001), and only vary in the level of product analysis and change-notification offered to customers.
If your lab were performing quality control or assurance in a regulated industry, you'd need to ensure that all of your reagents (and laboratory practices) are in compliance with such regulations. If you're in a research lab, these extra features may or may not add any value to your work, depending on the specific applications you have in mind. For most experimental applications, there's not likely much added value in being notified every time a manufacturer gets a different raw materials supplier since most LB ingredients like tryptone and yeast-extract are already complex (undefined) substances. Then again, if your research aim is to optimize growth for industrial fermentation reactors, that information might come in handy.

This answer only concerns Streptococcus pneumoniae.
The lab I worked in used  cheap ingredients, the same as everyone else claimed to in this field.
The only shocking thing I found was that the sterile media (already prepared) that I was given to use  for DNA extraction did not work (DNA would be fragmented). It turned out the media had been waiting in the incubator for a year. Made fresh media and all was well.
Todd-Hewitt Broth (THB) with 0.5% Yeast Extract (YE)
3.0 % (w/v)  of Bacto™ Todd Hewitt broth powder (Difco cat# 249240)
0.5 % (w/v) of Bacto™ yeast extract (Difco cat# 288620)


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Is it necessary to calculate lane normalisation factor when doing western blot data analysis?
I have recently done my first western blot and I am doing data analysis to quantify my blot. I have labelled my membrane against inactive GSK3 and active GSK3 which are phosphoproteins so I am using total GSK3 as an internal loading control. I have read some guides and handbooks  about western blot data analysis and I have seen that some of them calculate a lane normalisation factor to account for variations in signal intensities of the loading control. For all loading control bands in each lane, they divide by the loading control band with highest intensity to get the lane normalisation factor. Then for each band of the target protein of interest they divide by the lane normalisation factor to get the normalised intensity.
I was wondering in general with western blots is it good practice to calculate the lane normalisation factor when doing the data analysis? Any insights are appreciated.

Generally speaking, the proper way to quantify a western blot is to normalize to a loading control such as Actin or GAPDH. In this case it would be (pGSK3/Actin)/(GSK3/Actin) as total GSK3 is not a loading control. A loading control is to protein that is accepted as unvarying in concentration across multiple samples if the same protein amoun is used, and to my knowledge GSK3 would show more biological variability than accepted loading controls. This would be considered the best practice to compare protein concentrations across samples.
That being said, if all you care about is the ratio of active to total, pGSK3/Total GSK3 is probably sufficient to assess if your manipulation changed active/inactive:total ratios. For example, in autophagy assays you can measure autophagic flux by measuring LC3-I to LC3-II, and I've never seen them normalized to a loading control since its a ratiometric readout contained within a single sample.


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Can proteins be located on the surface of the mitochondria?
I am learning about the mitochondria and I know there are proteins present in the mitochondrial matrix such as SOD2, but I was wondering for a protein to be located on the surface of the mitochondria does it always have to be a membrane protein (e.g. inserted within or associated with the mitochondrial outer membrane) or can other proteins which are not membrane proteins be present on the mitochondrial surface?
Any insights are appreciated.

What kind of protein? Do antimitocondrial antibodies count? What function might it have, to be near but not integral to the mitochondrial surface? I'm not sure what you're asking.

I believe that one needs to think chemically about this. If a protein is present on the surface of a membrane, there has to be an interaction causing it to be there. By definition such a protein will be a membrane protein — either integral or peripheral. So are you asking whether there are peripheral outer membrane proteins?

Yes, integral membrane proteins interact with and form complexes with proteins that do not directly interact with the membrane.
Signal transduction often occurs when a soluble protein comes in from somewhere else and touches a membrane-bound protein.
Complexes often take the form of one anchored protein bound to additional proteins.
You can look at the full list of proteins associated with the mitochondrial membrane here, i bet if you go through them you will start to find ones that don't have a membrane-inserted domain.


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Do stem cells have no epigenome?
Till now I thought that embryonic stem cells have no epigenome as they are pluripotent. (I thought that since epigenome is what gives a cell its identity, no cellular identity means no epigenome) I saw something similar to this on this Wikipedia page. After fertilization, the paternal and maternal genomes are demethylated in order to erase their epigenetic signatures and acquire totipotency..
Other sources mention 'reset' in place of 'erase'.
This paper rather suggests that stem cells do have an epigenome. Specifically, genes associated with self-renewal are silenced, while cell-type-specific genes undergo transcriptional activation during differentiation..
I am not very literate in biology, please excuse me if I made a mistake.

So there are a couple of things to bear in mind.

pluripotent does not mean that all genes are active. It means that the stem cells have the ability to form different cell types. However, it still needs to keep the cellular programme of a neuron for example silent. So the epigenome is still present to keep other cell type programmes silent until there is a transition.

DNA methylation is not the only source of epigenetics. Active and inactive genes also correspond to particular post translational modifications on tails of histone proteins. In the cell, DNA is wrapped around histones to form what is known as chromatin.

Hope that is a starting point to answer your question


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Can apes have schizophrenia?
Many animals can become mentally ill, but I could not find any account of animals showing schizophrenia, not even apes. I could only find this, but it seems very vague.
Is there any evolutionary explanation of why only humans have schizophrenia?

Makes me wander how would you verify the animal has halucinations, delusions or disorganized thoughts.

@BagiM - You don't need to ask a Great Ape (or a mouse) if they're having hallucinations. If schizophrenia is genetic and the sites responsible for it are known, we need only to sequence DNA to know. If it is morphologic, we need only to do histological studies. And yes, both of these types of studies have been done on humans and great apes, looking specifically at schizophrenia as a uniquely human disease. I an not a neuroscientist, so will not comment on the findings, as they are kinda beyond my ability to judge competently. But a neuroscientist should be able to answer.

I really like this question (so +1), but I’m voting to close it because it might be a better fit on Psychology & Neuroscience.SE

@anongoodnurse: Re "If schizophrenia is genetic...", but is it purely genetic?  Or is it a case of the genetic variants being necessary but not sufficient?  In the same way that having certain genetic variants predisposes one to various cancers, but doesn't guarantee that you're going develop that cancer.

@jamesqf - (I hope you know what I'm going to respond...) Schizoprenia, like most mental illnesses showing a genetic component, is very likely multifactorial, with different factors having different weights/influences on expression. I do believe that the genetic component(s) is/are heavily weighted. My comment was to someone who doubted on the basis of asking an animal questions. A nuanced response would be kinda inappropriate to that user. I was pointing out that there is more to science than asking an animal if it's having, say, flight of ideas, or if their cat is giving them commands.

Like hearing voices? that's a characteristic symptom of one form of schizophrenia. or multiple personalities, which is the original meaning? you have to clarify. your question is a bit dull for effort at the moment. You can try this though https://scholar.google.com/scholar?hl=en&as_sdt=0%2C5&q=psychosis+chimpanzee&btnG= and https://scholar.google.com/scholar?hl=en&as_sdt=0%2C5&q=schizophrena+chimpanzee&btnG=


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Nomenclature of substrates for DNA synthesis
I have read in my school textbooks that both deoxyribonucleoside triphosphate and deoxynucleotide triphosphate are used in DNA Replication as substrates.
However, it is unclear to me whether the terms refer to the same molecule as one uses the term nucleotide and the other uses nucleoside. What confuses me is that nucleoside is a compound with a purine or pyrimidine base and a sugar and a nucleotide is the same but with the inclusion of a phosphate group.
Is it possible that the “deoxynucleotide triphosphate“ is used in artificial DNA replication as in PCR, and “deoxyribonucleoside triphosphate“ is the natural substrate for the replication of DNA within cells?

The short answer is that one can be almost certain (as certain as possible without seeing your textbooks) that both of these terms refer to the same thing: 2′-deoxyribonucleoside triphosphates. There is no difference in the substrate requirements for synthesizing DNA in vivo or in vitro (as with PCR), but it would be a worthwhile exercise for you to consider what would happen if you added 2′3′-di-deoxyribonucleoside triphosphates to a reaction mixture. If the question is not judged by others to be off-topic I may elaborate on the nomenclature and what parts are important when I have a moment.

Thank you so much sir! We have been taught about ddNTPs, if that is what you are suggesting. I believe that they are used in Sanger's method for determining DNA sequence as they lack an -OH group at 3' which is important for chain elongation and without this 3'OH the chain synthesis is terminated. And I would be very grateful to you if you do get a chance to elaborate on the nomenclature. Much obliged.

Summary
The systematic chemical names of many important biological molecules are too long to write routinely in full, and in any case were preceded historically by abbreviated forms which generally stressed the distinguishing features of importance to those studying their metabolism. Thus, as the poster points out, a molecule such as dATP (image below) has a base-(deoxy)sugar-phosphate(s) structure, and can be termed a deoxynucleotide. Adding triphosphate provides the additional information that there is more than one phosphate. A base-sugar(deoxyribose) combination is, as the poster states, a deoxynucleoside, but by extending this name to deoxyribonucleoside triphosphate, one indicates the addition of phosphates, so that one is, by definition, describing a deoxynucleotide. The reference to ribonucleoside indicates that the sugar is ribose, a specification lacking from the other definition. However, in practice these two terms are used to describe the same entity, one which might a little more completely describe as 2′-deoxyribonucleoside triphosphate (to indicate the position of the reductive change). This answer attempts to explain with precision the aspects of the nomenclature that are likely to be important to the biochemist or molecular biologist.
Disclaimer
The original question might be considered not to represent a biological problem, and one might argue for closing it on that account. However because of the very nature of trivial nomenclature, it is difficult for those without experience of the usage in a field to find the answer. In answering it I have tried to broaden the scope to include the aspects of structure that I feel are most likely to be to biological students.
Answer
It is convenient to start with ribonucleoside triphosphates (rather a deoxy- ribonucleoside triphosphates), and I have chosen a specific example in the molecule that is generally known as ATP:

Alternative names for this ribonucleoside triphosphate, in decreasing order of complexity, include:

[[(2R,3S,4R,5R)-5-(6-aminopurin-9-yl)-3,4-dihydroxyoxolan-2-yl]methoxy-hydroxyphosphoryl] phosphono hydrogen phosphate
9-β-D-ribofuranosyl adenine 5′-triphosphate
adenosine 5′-triphosphate
adenosine triphosphate
ATP

Of interest to us are the three components of the molecule: a purine base (adenine), a sugar (ribose), and three linked phosphates. This is not reflected in the chemical nomenclature (1), so let us focus on the, albeit rarely used, biochemical nomenclature (2).

The ‘9’ specifies the position of the adenine ring that is linked to the sugar. This is not of particular interest in the context of the question.  (The numbering of the ring is, however, relevant to describing differences between or modifications  of the purine and pyrimidine bases1.)
The D-ribofuranosyl is a form of ribose (see below). The ‘D’ indicates that it is D-ribose enantiomer rather than the alternative, L-ribose. However, as all natural sugars are ‘D’ this is generally omitted (although discussions of why all natural sugars are ‘D’ abound2).
Ribofuranosyl indicates that the ribose is in the five-membered hemiacetal ring form (rather than the straight chain or the six-membered pyranose ring), but this is always the case in nucleosides, and so is not of particular interest.
The 5′ specifies the position of the sugar ring to which the phosphates are attached, the prime (′) indicating that the numbering is for the sugar ring (not that of the base). This is important, as it is relevant to the linkage in the (asymmetric) phosphodiester bond made to the 3′-OH:

The image above is for DNA, but before addressing the nomenclature there, we would point out:

The common α, β, γ designation of the three bond phospho-ester bonds. This is very relevant to the formation of polynucleotides (DNA and RNA) from their substrates, and in general to the utilization of the free energy the hydrolysis of these bonds for other processes3.

Finally we come to a structure of interest to the poster, dATP:

In the designation, 2′-deoxyadenosine 5′-triphosphate, the 2′-deoxy indicates that it is the 2 position in the ribose ring (′) that has had the oxygen removed (deoxy), i.e. OH is reduced to H. Although common knowledge, it is important that the student new to the subject understands this, as if it were the 3′ position that was reduced, it could not form the phosphodiester bonds of DNA. And it is relevant in many other contexts. For example the chain-termination method of DNA sequencing uses 2′,3′-dideoxy nucleotides to inhibit such bonds forming4, the single difference between ribose and deoxyribose allows us to understand the profound affect on the susceptibility of RNA to alkaline hydrolysis in nature5, and certain RNA molecules are hydroxy-methylated at this position.

Related information in SE Biology answers
1 Topical example related to base modification
2 Speculation on the origin of biological homochirality — not SE Biology
3 Why does ATP sometimes produce ADP and sometimes AMP?
4 Sanger sequencing by chain termination
5 Difference in stability of DNA and RNA


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What genes are required to make E. coli photosynthetic?
"All" of the genes for bacterial photosynthesis were discovered in a gene cluster almost 40 years ago. Marrs, J.Bact.  What more is needed to make E. coli photosynthetic?

Related post https://biology.stackexchange.com/q/17676/3340

I think the crux of the answer is only hinted at in WYSIWYG's answer. The crux of photosynthesis is partitioning coupled reactions.

Photosynthesizing plant cells have sub-cellular structures called chloroplasts; their membranes are especially important for the compartmentalization of different functions and chemical processes for the entirety of photosynthesis to even be possible. Partitioning is difficult to achieve in E. coli, and that to me seems a necessity for achieving true heterologous, truly ectopic photosynthesis... Even photosynthesizing cyanobacteria (photosynthesizing prokaryotes) possess thylakoids, membrane-bound compartments where the magic happens. Photosynthesis though can certainly proceed and be made more efficient in single cells (example).
That leaves us with a simple answer: engineering E. coli with thylakoid-like structures is not really practicable. Why not simple use cyanobacteria instead?
Further reading: review on single cell photosynthesis in single cells.

A very basic kind of photosynthesis can be realized in E.coli by expressing a rhodopsin (Kim et al., 2017). Rhodopsins are membrane proteins that can pump ions (including protons) across the membrane and generate a transmembrane potential. When coupled to ATP synthase, rhodopsins can cause ATP generation in the presence of light. In an earlier work, Hara et al., (2013) had expressed a rhodopsin in mammalian mitichondria to make them synthesize ATP in response to light exposure.
This is not the kind of energy efficient photosynthesis that we see in cyanobacteria and plants. The group of Ron Milo have engineered Calvin-Benson cycle (carbon fixation) in E.coli, and optimized carbon fixation using adaptive laboratory evolution (Antonovsky et al., 2017; Gleizer et al., 2019).
Engineering the entire photosystem in E.coli would be a difficult task because a functional photosystem does not merely involve expression of the pigments but also their 3D assembly into functional sub-cellular "organelle" like structures.

Has either a simple anoxygenic reaction center or bacteriorhodopsin been expressed in E. coli in a functional manner so that the organism can energize its membrane via light?

@Youvan Check out the first reference. It also cites previous work on this topic as well.


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Which terminology is better: Heterologous expression system or heterologous cell model?
I am reading a journal paper and I have read that the authors expressed the extracellular domain of the protein NCAM2 which is normally found in the brain in CHO cells. I know that this is an example of heterologous expression.
However, when referring to these CHO cells which are overexpressing NCAM2 I am wondering whether the terminology 'heterologous expression system' is more appropriate compared to 'heterologous cell model'?
Any insights are appreciated.

I would suggest that the preferable term depends on whether you are talking about the cells or talking about the modifications.
In other words, I would refer to the NCAM2 expression package as the "heterologous expression system", which is then integrated into the CHO platform in order to create a "heterologous cell model."
My reason for this suggestion is that the expression package is the core construct without which there is no expression, and that this package might potentially be transferred into other cell lines to construct other cellular models.


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Alternative dyes for Gram staining
In the Gram stain, is there any replacement for primary stain, secondary stain, decolourizer and mordant? What results will the replacements produce?
I found that crystal violet can be replaced with methylene blue but in that case, what will the secondary stain, mordant etc. be? Please also explain what result we might get.

As you may have realized, crystal violet can be replaced by a lot of dyes since the ethanol will wash out the dye from the gram negative cells. Methylene blue is a nice choice - malachite green may also work.
A weak concentration of hydrochloric acid (3%) is also a possible replacement for the ethanol. A 3% HCl solution is used in the acid-fast stain but it may be too strong for non-acid fast cells. It would be worth testing it out and trying it to see if it could replace the ethanol step.

Welcome to Biology.SE! This looks like it might be a good answer, but answers are much more likely to receive a favorable response if you include supporting references (primary literature is best). Without that support, your answer is indistinguishable from opinion to a non-expert — it also helps users by providing resources for further learning. ——— A good example of how to format references. ——— You may also want to take the [tour] and then consult the [help] pages for additional advice on [Answer] effectively on this site. Thank you!


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What are the advantages of heterologous expression systems?
I am doing research in a lab and I am relatively new to wet lab work. Many experiments that people are doing in my lab include overexpressing a protein of interest from another species (e.g. mouse) into CHO cells via transfection. I have learnt that these are examples of heterologous expression systems.
I was wondering in general, what are the advantages of heterologous expression systems? I have read that you can use them to study the function of a protein in a well-defined, controllable environment. However, I was wondering what other benefits/uses are there to these kinds of experiments?
Any insights are appreciated.

It takes significant resources to establish a cell line. More efficient to use existing lines.

Often cell lines used for expression grow faster. This is why so many proteins are expressed in E coli - they only take a day to grow vs the week or two for mammalian cells, meaning you get a much faster learning iteration in your research.

To add on @ user47696’s answer, heterologous expression systems also enable large-scale production of recombinant proteins, particularly in the E. coli and the yeast systems. Foreign hosts may also provide a system that is simpler than the natural sources for the studies on proteins' functions.


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In western blots, do loading controls have to be proteins that are in equal amounts in each lane?
I have recently done a Western Blot and I am doing data analysis on my blots. I am studying the protein GSK3, which is a phosphoprotein.
I have labelled my membrane against active GSK3 and inactive GSK3. I have also labelled the membrane against total GSK3. I have used Ponceau S as a loading control. From my Ponceau S staining I have seen that there are variations in the amounts of protein loaded per lane. So there are variations in the amounts of total GSK3 per lane (the intensities of the bands are not all equal).
I am wondering if the band intensities of total GSK3 are not consistent among lanes, in this case would using total GSK3 be a good loading control to normalize my data on? I have read for the case of phosphoproteins, you can normalise to the total protein (using an antibody that recognises all forms of the protein of interest).
I have read that loading controls are endogenous proteins that are unaffected by experimental conditions and used as an indicator of sample concentration. But if the amounts to total GSK3 vary between the lanes, how can this be a good loading control? Similarly, why do many researchers use Ponceau S as a loading control, if the band intensities among lanes vary (if you don't load equal amounts of sample)?
In other words, for a protein to be a loading control, does it have to be in equal amounts for all your samples/lanes?
Any insights are appreciated.

Can you clarify what you are loading? Purified GSK3, total protein, or something else.

I am loading 20 micrograms of brain lysates per lane (total protein that is present).

Controls do not need to loaded in equal amounts per say, but they do need to be expressed relatively equally between lanes. They need equal expression, such that controls can be used to normalize values of the experimental proteins.
It is common to use housekeeping genes, such as beta actin, which are usually expressed equally between treatments to control for loading.  Important to ensure treatments don't affect those proteins.
For examples and rationale, see:
https://www.abcam.com/primary-antibodies/loading-control-guide
https://www.bio-rad-antibodies.com/western-blot-loading-controls-antibodies.html
https://www.labome.com/method/Loading-Controls-for-Western-Blots.html


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How broad are the conclusions that one can make from a heterologous expression experiment?
I am studying whether a neural protein regulates the activity of the protein GSK3. I am using heterologous expression systems where I am overexpressing the neural protein in CHO cells (via transfection) to see if it affects the levels of GSK3 in transfected cells vs controls. I am interested in seeing whether the neural protein regulates the activity of GSK3, and I have found that the levels of GSK3 are higher in the transfected cells vs control. But as I am using CHO cells as a model system, how broad are the conclusions that I can draw from this experiment? For example, will I be able to generalise what I found to neurons, or hypothesise that the same thing that I observed in this experiment occurs in neurons? Any insights are appreciated.

Hi, @ceno980: I've noticed you asking a number of questions that would be well-suited to the proposed Synthetic Biology SE site. Want to join the proposal?

GSK3 is interesting because as a kinase, it is itself regulated by its phosphorylation status (particularly by Akt in neurons). Therefore, more protein does not necessarily mean more activity. For example from E13.5 to E15.5 GSK3 levels do not change but activity steadily increases. Hence if you increase GSK3 levels in neurons, it may not mean that downstream effects would change substantially, so I think although promising the in vivo effects may be hard to deduce from this experiment. If you see similar effects in in vitro neurons, you can be a bit more confident of potentially seeing similar effects with in utero electroporation in vivo


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Please identify this syrphid (hoverfly)
Last summer I took these pictures of a member of the Syrphidae on my balcony (Switzerland, Geneva area, Alt. ~380m, June 2020 @19:14)
Pictures:

I hesitate between three species but it could also be another one:

Scaeva dignota
Scaeva selenitica
Scaeva pyrastri

Because the pale marks on tergites 3 and 4 seems parallel to the anterior margins of the tergites, I would eliminate the third one, especially because the marks are also more yellowish than white.
For the two other species, I am not sure because I checked many pictures on the internet, but some are wrongly tagged (google images results).
Any entomologist can confirm the exact species of this hoverfly?

Awsome pictures, we have them in Austria too ;) Personally I tend to the 'Scaeva selenitica', but that's just my uneducated guess..


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Differences between comatosed states and brain death
What are the neurological, physical, etc. differences between someone who is in a coma (one that is not medically-induced) and someone who is colloquially deemed "brain dead" (being physically kept alive by machines)? Is there a certain minimal brain activity that is present in a comatosed state that is not present in brain death?
I found a decent definition here which defines the difference as:

Brain death: Irreversible cessation of all functions of the entire brain, including the brain stem. A person who is brain dead is dead, with no chance of revival
Coma: A state of profound unresponsiveness as a result of severe illness or brain injury

I can find other similar, vague definitions in a few other places.
I am wondering what is different about the neuronal structures or electrical activity between brain death and coma. Is there simply no (zero) neural/electrical activity with brain death, whereas there is with coma? What would be physiologically different with brain death that would prohibit such neuro-electrical activity?

Why the DVs? The question is on topic, is not a dupe and shows preliminary research. I had asked this on MedicalSciences.SE and they told me to ask it here instead.

I suspect the issue is that you haven't shown any research or attempt at answering your own question. 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. Additionally, the answer is kind of obvious - the difference between someone who is comatose and one who is brain dead is that the comatose person still has brain activity, while the brain-dead person does not. You need to specify what else, if anything, you want to know beyond that.

Thanks @MattDMo (+1) - please see my edits. Online definitions (like the one provided) are all vague and seem to target edifying the laymen.  I am wondering from a physiological perspective: what would cause brain death to the point where there is no neuro-electrical activity possible or observed across the brain? But, that does not inhibit such activity in an individual who is only comatosed?

I'd recommend reading a review paper on disorders of consciousness, which cover the spectrum of minimally conscious states, vegetative states, and coma. Brain death is outside the context of disorders of consciousness, but is often discussed for comparison. Nico Schiff is one expert in the area; I've attached a reference to a review he coauthored at the bottom and I'll draw mostly from that review in this answer.
Schiff and Fins write, with some bolded emphasis by me:

At present, brain death is diagnosed if clinical evidence of the complete loss of brainstem function is present at the bedside and the patient demonstrates a failure of ventilatory drive in the setting of documented hypercapnia (significant elevation of blood partial pressure of carbon dioxide over a chosen threshold) — the ‘apnea test’. Fulfillment of these criteria should invariably associate with an ‘empty skull’: no sign of any metabolic activity or blood flow measured by positron emission tomography, a ground truth linking the biological model to the clinical assessments

The first two statements are most relevant clinically; PET is not commonly used except in research and rare cases of uncertainty. Biologically, brain death is when the brain is...dead. There is no recoverable neural function. The signs are a lack of any brainstem reflexes, including lack of respiratory drive. Additionally, other potential causes of these symptoms (such as pharmacological causes) must be ruled out.

Coma is a state of unconsciousness, but some brain functions are preserved. Again from Schiff and Fins:

Both comatose and vegetative state patients are unresponsive to environmental stimuli, although reflex movements may be present; in neither condition are goal-directed behaviors initiated. Comatose patients lack state variations and typically remain in a closed-eyes state that is unchanging even when presented with the most vigorous stimulation.

Comatose patients do not meet the criteria for brain death described above, but do have a complete inability to respond to stimuli and do not make purposeful movements besides reflexes. Schiff and Fins (and others) compare this state to deep anesthesia. Though your question posits that there should be some distinction between medically-induced and one that is not, there is really no functional difference. The possibility of recovery is tricky to assess, but generally patients who spend a long time in coma have a worse prognosis than those that do.
The clinical approach is different. A brain dead patient is dead. Nothing can be done to recover function; potential for organ donation can be considered, but there is nothing medically to do for the individual. For a comatose patient, it's more complex, and decisions need to be made regarding the level of medical support to provide given expected prognosis in both probability and extent of recovery.

In brain death, the neurons are dead. The lack of electrical activity is not because they are in some quiescent state, it's because they are dead. With coma, it's less clear except that there are signs that some neural function remains. Coma is really a clinical state, not a biological one. It's not always possible to determine the level of irreversible damage in a comatose patient, nor the extent of possible recovery.

Schiff, N. D., & Fins, J. J. (2016). Brain death and disorders of consciousness. Current biology, 26(13), R572-R576.


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Immunological factors for the cause of headaches following SAR-CoV-2/COVID vaccination
It is generally-accepted that headaches are a common side effect from receiving the COVID vaccine.

Vaccine recipients with pre-existing immunity experience systemic side effects with a significantly higher frequency than antibody naïve vaccines (e.g., fatigue, headache, chills, fever, muscle or join pains, in order of decreasing frequency, P < 0.001 for all listed symptoms, Fisher’s exact test, two-sided).

— Florian Krammer et al.: "Robust spike antibody responses and increased reactogenicity in seropositive individuals after a single dose of SARS-CoV-2 mRNA vaccine" doi: https://doi.org/10.1101/2021.01.29.21250653
I am trying to understand why. What theories or known biological, immunological or otherwise physiological mechanisms would cause recipients of the vaccine to report headaches as a common side effect?

Headaches are a side effect of both vaccinations and infections.  Have a look at the effects of cytokines, particularly Interferons.

Headache is one of the various side-effects of vaccination. Typically, a person will have pain in the site of injection, muscles throughout your body and head.
The immunological reason is the network of our immune system. Whenever a foreign antigen (vaccine) is detected by our immune system, antigen-presenting-cells (macrophages, dendritic cells) takes the antigen to nearest lymph nodes where it is presented to the adaptive immune system. Due to this invasion, strong immune response starts which leads to the pain.

Image Source: wikipedia.org/wiki/Lymphatic_system

Thanks for the amazing answer @Riq (I would upvote you +1 but I don't have enough rep!) -- one quick followup question for you, if you don't mind. It sounds like you're saying that when cells, such as macrophages, present lymph nodes with antigens (from the vaccine) this initiates an immune system response, which you say leads to "pain". But why headaches? Why is this pain so specific to the head? Thanks for any clarification!

BTW I am not the one who gave you a downvote! I am happy with your answer so far!

@IamYourFaja You can accept the answer if you think it answered your question.

Yes thanks again @Riq, do you have any ideas regarding my followup question above ("But why headaches?...") Perhaps something to do with the cytokines and interferons that Polypine mentioned?

@IamYourFaja yeah, cytokines are essentially the communicator of our immune system. They help in cell signaling and involved in various side effects of our immune system. Hope this clarifies your follow-up question.


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What are the units of the band intensities in a western blot image?
I have done a Western blot and I am measuring the band intensities in Image J. Using the rectangle tool, I have outlined the band and then calculated the area and the mean intensity of the band. I have then calculated the total intensity of the band by multiplying the area of the band with the mean intensity. I was wondering, what are the units of the band intensities? Are they in arbitrary units? Any insights are appreciated.

The units in your case are arbitrary units.
For instance, you may have taken a photo of the bands with a camera that saved a gray-scale file with an 8-bit depth. That means that the brightness values of each pixel have 256 gradations (or 2^8 values of brightness possible between minimum/blank and maximum/saturated). The program only takes the pixel brightness value anywhere between black (0) and fully white (256) and simply performs the calculations so. There is no real connection between actual band intensity and the output from ImageJ or any software.
Alternatively, you can also have relative units. Say you have a reference: a positive or a negative control. If your reference has a brightness of 10, and your band has a brightness of 7, you could express the band intensity as "0.7 relative units"; here you have to specify your reference, i.e. what you are calibrating your relative units to, hence the term relative.


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Why do cells in meristematic tissues lack vacuoles?
Cells located in meristematic cells in plants lack vacuoles. However, this contradicts with the fact that plant cells have large vacuoles to store water and keep the plant in an upright position. I would also like to add that meristematic tissues are generally found in roots and stems of plants. Finally, I would like to ask why do meristems lack vacuoles?
Sources

Turgor pressure exerted by the vacuole is also essential in supporting plants in an upright position: Wikipedia

Apical mertisem is present at the growing tips of stems and roots and increases the length of the stem and the root. The girth of the stem or root increases due to lateral mertisem (cambium): Science for Class 9 by National Council of Educational Research & Training, page number 69

Cells of meristematic tissue are very active, they have dense cytoplasm, thin cellulose walls and prominent nuclei. They lack vacuoles: Science for Class 9 by National Council of Educational Research & Training, page number 70

Please do not edit the title to "answered", please accept the answer.

@Chris Well, I need to wait sixteen more hours to do that. Thank you!

I haven't looked at the times :-) It would be good if you could do it, then it is marked as answered and accepted on the starting page.

Meristematic tissues or meristems contain cells which can rapidly divide or reproduce and is responsible for the overall growth of the plant. Apical meristems and lateral meristems are found in the roots and stems of the plants. Please note that the whole stem or root is not laded with these tissues. Plant cells usually have a large central vacuole which applies turgor pressure to keep the plant upright, as mentioned in one of your sources.
The tips do not require to be very stiff and straight. So, a central vacuole is just not required. Also, meristematic cells grow various organelles as they mature.


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Does the Miller-Urey experiment explain the origin of life?
According to the Miller-Urey experiment, the early earth atmosphere could have supported the formation of amino acids - and the experiment is hailed as being important in the explanation for the origin of life. However, I do not understand how life could have started from amino acids (or proteins) which do not self replicate. Certain types of RNA do self-replicate, and it is plausible how life could have evolved from there.
Does it matter if amino acids came before RNA if they did? Is it correct to assume that in the RNA world, life somehow learnt how to use amino acids?

"the experiment is hailed as being important in the explanation for the origin of life" Please consult sources that discuss this and note exactly they say, i.e. why they think this experiment is important. Any source that does not say so is of no use to you, but I am sure you will find respectable sources that provide reasons. If you still want to know the significance of the Miller-Urey experiment, update your question explaining what you do not understand. But do research and pose precise questions on a topic that can only have arguments for answers.

Disagree with David. This is a common question and does not need reducing.

This is a common question. I think the experiment and its conclusions are often misunderstood.
Originally, the null hypothesis was that the ingredients for life could not have come about spontaneously, from inert, inorganic molecules. There was no evidence of strictly physical, non-biological processes being able to produce compounds that were necessary for the proteinaceous life as we know it. In other words, abiogenesis was therefore not supported by any line of positive evidence... until the Miller-Urey experiment. The experiment showed that the ingredients could arise through plausible, natural conditions prior to the existence of life! It bridges the gap between inert, inorganic, non-biological, and the rich soup of complex molecules that would - arguably - be necessary for anything like abiogenesis to even be a consideration based on empirical observation.
Of course the presence of amino acids is not evidence for the abiogenic origin of life. Life is not protein nor amino acid alone. But demonstrating that complex biological ingredient chemistry occurs spontaneously in large abundance - that sounds like a great take-off point for an abiogenic origin of life! That's really all the experiment achieved. In its historical context, it is a very impressive discovery, but it is certainly not a complete explanation, merely something that makes myriad biochemical explanations possible (and perhaps even plausible!). Perhaps you can now better appreciate why it excited and continues to excite biochemists working on trying to understand the chemical origins of life. The Miller-Urey experiment is foundational.
As for the transition from RNA to an RNA-protein origin of life, I quote briefly from another answer elsewhere here:

Regarding the transition from RNA-only to RNA-protein world, peptides
function as cofactors for some ribozymes. Amino acids and peptides are
known to have existed in the prebiotic environment and have been found
in space (glycine has been found in comets, along with other 70 amino
acids).

The actual problem with this experiment is that the environmental conditions implied in it might have never existed on Earth - at least, this is what the geologists say.

The discussion about the historical validity of the conditions assumed in the experiment has been ongoing and rich and full of disagreement, even among experts; similar experiments have been performed with many adjustments since. It's worth taking a look into it if interested!

This is reaching back several decades, but has the Miller Urey experiment ever been replicated? I thougt not... am I mistaken? I'd love to know.

The Miller-Urey experiment does not explain the origin of life. It just propose a plausible way to develop life from organic matter, as part of the abiogenesis model of the origin of life. A better approach to understand the theories of life origin is looking for a context.
We already know, by the cellular theory, that cell are the main component of organisms and that just cells could originate others cells. So, the research is focused on finding a way to create at least one cell. From knowed compounds, cells could be modeled as a set of autoregulated biomolecules. Just two of them are used explicitly for replication: nucleic acid and proteins. If you think about this two components, the Miller-Urey experiment just propose a way to develop simple proteins from aminoacids, and aminoacids from inorganic matter. Amazing, but not enough for explaining the complexity of actual proteins and enzymes.
But there are other theories and facts that works well with this experiment. Some non-translated olygopeptides have intracellular functions. The PAHs world theory proposes a way to develop nitrogenated bases from aromatic hydrocarbons. This bases are precursors for RNAs and DNAs. The RNAs world hypothesis establish a way to develop life from RNA and not DNA or proteins. The phospholipid bilayer have some common patterns with micelles and all this mechanisms would be part of the Last Universal Common Ancestor (LUCA). But there is no evidence of the relationships between this theories and maybe there are just too simple for explaining life.
Maybe the first cell had come from the space, where a pre-term version the krebs cycle have been demostrated to work. We could think on it as "biogenesis" until "abiogenesis" could be demonstrated.

"first cell from space" does not solve the problem it just side steps it, also you may be interested is some of the more recent work on RNA synthesis. https://pubmed.ncbi.nlm.nih.gov/33547911/


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Enzyme inhibitor leads to higher turnover rate?
I'm currently working on a project where I have to deal with enzyme inhibition.
The purified enzyme shows a good substrate turnover. When I try to inhibit it with different inhibitors described in literature, however, I receive quite awkward results.
I have now tested 3 different inhibitor compounds. One showed good results (it reduced the substrate turnover), while the other two compounds instead led to increased substrate turnover?!
I know it sounds ridiculous, but has someone experienced something like this before? The measurements of the "working" and "not working" compounds take place on the same plate.
How can it be that widely described inhibitors lead to such results?

This is very vague (purity of the enzyme, etc.) You might be messing wit the rate limiting step somehow. But my question is, why aren't you asking someone senior in the lab? Is this a homework question?

Adding on to anongoodnurse, you say the rates change but...by how much? What's the variance in your measurements? How many replications? How are replications performed? In what order did you do the experiments and how were the reagents handled during the process? What other conditions are you maintaining constant?

What do you mean by a “Signal increase”? This is not a term I have ever encountered in enzymology. Please reword your question using standard enzymological terms. Running your question through an English language spelling checker is also recommended. Modern software and operating systems make this easy. I routinely do checks in French, German and Italian.

@David The language was a bit rough, but I found the meaning pretty clear, as well as the starting points for debugging. I have now cleaned it up and hope others will retract their close votes.

@jakebeal I am afraid this question is still completely unclear, and until the poster uses the precise scientific terminology to describe what he observes — together with detail on the so-called "inhibitor compounds" the question cannot be answered. Nor does it deserve to be as the poster has not responded to this basic request to use enzymological terms, leaving one to wonder whether he understands the basics of enzyme measurement and quantitation. The term "substrate turnover" typifies this problem. I spell this out in my next comment.

One measures "the rate of enzyme reactions", which when expressed in standard terms (e.g. nmol substrate converted per min) is referred to as activity. To convert this to meaningful information for enzyme or effector (inhibitor or activator) one determines activity over a range of substrate concentrations to calculate the Km and Vmax (from which the  turnover number can be determined for pure enzymes). The type of inhibition can be deduced from the effect on these parameters and the kinetics (MM, allosteric etc.). You need this information to make sense of your results.

@user338907 — If your remark was directed at me, my point was that there are standard methods of studying and assaying enzymes and the effects of inhibitors and the poster should describe which if any he has used. I imagine he has done none of these, so nit picking my comment is unhelpful. I would make an observation about the terminology remaining in the edited question — "substrate turnover". In general turnover implies that something is used and then replenished, as in protein turnover. Turnover number relates to the active site cycling between occupancies. Substrate does not turn over.

While it's difficult to know just what is going wrong in somebody else's laboratory, apparently paradoxical effects like these are unfortunately common in laboratory work.  Assuming that these are indeed well-established inhibitors that really should be working, let's consider why they might not be in your hands.
The mechanisms of molecular interaction are complex enough that there are many cases where a substance that acts as an activator in one condition can act as an inhibitor in another condition. Another way that you could see such an effect is if your purified system is not actually as good as you think and the problem is being partially relieved by one of the inhibitors, e.g., imperfectly purified and addition of the inhibitor neutralizes some of the imperfections.
Not knowing the details of the compounds you are working with, it is not possible to guess which of the possibilities might be at work. I would suggest, however, that you begin by:

Comparing your quantitative turnover rate with the purified enzyme to the predicted rate from the literature to see if your baseline system is actually working correctly.
Make sure your system is also (not) processing substrate at the appropriate low rate when the enzyme is absent.
Checking for potential differences between your method and the method that you are trying to adapt from the literature, which might give clues to the condition differences that may be impacting your results.

@user338907 Indeed: so many assays go wrong because there isn't sufficient use of process controls or because somebody only checked controls for the qualitative and not quantitative behavior. Your comment reminded me that the OP also didn't mention a negative process control, so I've added that as well, which would detect something like the case you mention.

There's also the factor of error - OP didn't mention the statistics they did on the data they generated, but it is highly likely that the curves are within "no significant difference" of each-other.


text:
What makes the neurons of the auditory nerve with such high-speed?
What makes the neurons of the auditory nerve with such high-speed? So the most labile are the fibers of the auditory nerve, in which the frequency of the generation of PD reaches 1000 Hz, while for skeletal muscles, it corresponds to 200 pulp / s, for smooth - 10-20 imp / s.  Motor nervous ending can be transferred to the skeletal muscle and at all no more than 100-150 excitations in 1 s

By "high-speed", are you referring to propagation speed or frequency? It sounds like you are talking about frequency rather than speed.

It's a very good question, but there are grammar mistakes and lack of precision and context. Perhaps you can rewrite it for a general audience?

There seems to be a verb missing from the title and first sentence of this question (between "nerve" and "with"). Not being a neurophysiologist I can only fire a guess. But I may be wrong.

High Speed vs High Frequency
For starters, what you are describing is a high frequency signal, not necessarily a high speed signal.  High speed signals are caused by myelination (a sheath of fat deposits that encases neuron fibers), and by minimizing the number of actual dendrite/Axon junctions you need to pass through to get from point A to point B.  In this respect, the nerves responsible for muscle control tend to be very long and have similar myelination to the cells in your ear; so, muscle control neurons would if anything transfer a signal faster than the more densely packed neurons in your ear.
Now to Answer the actual question: Why the Higher Frequency?
No one nerve in your ear actually needs to transmit a signal at a higher frequency than those nerves used by muscles.   Instead you have a number of different nerves detecting for different audio frequencies, and your brain then receives a combination of signals that tell it that various Audio Frequencies are being detected 1.  Each of the hairs in your inner ear tapers from the base to the apex such that the apex begins being stimulated by low frequencies, but the base only begins being stimulated by higher frequencies 2.
Each auditory nerve fiber is actually a cluster of nerves that work together at different frequencies to create a more complex signal. This results in a number of parallel signals that is interpreted as a complex (high frequency) signal.
Here is a simple diagram showing how low frequency signals can merge to make a higher apparent frequency.

Some animals can respond to sound in the microsecond range. They think that it's by using big groups of neurons working at the same time. Indeed, 1 cubic millimetre can contain 50,000 neurons / many receptors.
ref:https://journals.plos.org/plosbiology/article?id=10.1371/journal.pbio.2006422

Echolocating bats are powerful models for investigating the underlying mechanisms of auditory temporal processing, as they show microsecond precision in discriminating the timing of acoustic events. However, the neural basis for microsecond auditory discrimination in bats has eluded researchers for decades. Combining extracellular recordings in the midbrain inferior colliculus (IC) and mathematical modeling, we show that microsecond precision in registering stimulus events emerges from synchronous neural firing, revealed through low-latency variability of stimulus-evoked extracellular field potentials (EFPs, 200–600 Hz). The temporal precision of the EFP increases with the number of neurons firing in synchrony. Moreover, there is a functional relationship between the temporal precision of the EFP and the spectrotemporal features of the echolocation calls. In addition, EFP can measure the time difference of simulated echolocation call–echo pairs with microsecond precision. We propose that synchronous firing of populations of neurons operates in diverse species to support temporal analysis for auditory localization and complex sound processing.

If you search for "auditory microsecond" that's the measure in which the science is studied, so you will find lots of results there.


text:
How would you use readthrough as part of a genetic circuit design?
Traditionally, in synthetic biology, researchers tried to avoid some transcription phenomena (like roadblocking of tandem promoters or readthrough of weak terminators) since they are not in line with the discipline's strive to design modular, digital genetic circuits. However, some researchers have started using these "problems" as part of the intended design (see Bordoy et al., for example).
How do you think transcriptional readthrough could be used as part of a genetic circuit design? When would you want to have a readthrough? What kind of logic or function would this emulate?

Readthrough has already been used to implement transcriptional NOR gates, wherein tandem input promoters express a repressor that represses an output promoter. (First time I recall here: 21150903, used widely here: 27034378, roadblocking modeled here: 32141239.) An advantage of this design is that the repressor need only be encoded in DNA once, which can significantly reduce the DNA footprint of the design and the likelihood of recombination if the repressor's DNA sequence is long (e.g., protein-based repressors). OR gates have similarly been implemented as two tandem promoters expressing a common output gene. (An alternative NOR design—which is common with repressors with short DNA sequences (e.g., sgRNAs)—is to duplicate the repressor and express it separately from both input promoters. This approach circumvents the roadblocking and input asymmetry that can occur with tandem promoters.)
Readthrough has also been used with antisense promoters to tune gene expression (see 26769567). In these designs, an input promoter must read through a downstream antisense promoter, and the strength of the antisense promoter can be used to tune the response of the input promoter.
Another application that comes to mind is encoding the relative ratio of expression of multiple genes by separating them with weak terminators (e.g., to achieve X-fold expression of the first gene relative to the second). This begins to evoke an operon, as jakebeal notes. The closest example I can think of comes from this paper: 23727987, although they actually used readthrough to address the inverse problem of characterizing terminator strength.

I think this is an interesting proposal to try to turn a problem into a benefit.
What you are considering is conceptually similar to the organization of an operon, in which multiple genes are controlled by a single promoter, with multiple ribosome binding site (RBS) entry points for translation, and read-through can certainly occur in these cases too. Structurally, this would be the same behavior, except that the entry points are promoters instead of RBSs.
At a logical level, in effect the second promoter ends up implementing one of two functions:

If it is positively regulated, then it would be an OR, activated either by its own transcription factor or read-through from the 5' sequence.
If it is negatively regulated, then it would a NOT IMPLY, activated by the 5' sequence, but dominated by its own repression, since steric repression mechanisms will tend to block the read-through as well.

There are lots of potential concerns regarding signal strength of the read-through, but if one wanted to make use of this mechanism there is no reason to think they would not be susceptible to engineering.

Exactly. So how stochastic is the readthrough? Is there anything analogous to electrical engineering? How/when it is used in electrical engineering? Is there a logical equivalent for transcriptional readthrough?

I think the closest analog would be wired logic gates, though that's not asymmetric like read-through is. I wouldn't worry too much about the stochasticity, though: the promoter is stochastic too, so one would just want to appropriately match their levels.


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Dissection techniques for fruits and seeds
I'm doing some amateur botany, and I was keying some plants in the Brassicaceae (Cruciferae) family, which requires detailed information about the seeds of the plant. My question is, in a seed, how can one identify features such as:

Cotyledon(s)
Radicle
Endosperm (when present)
Accumbency/incumbency of the cotyledons with respect to the radicle (in dicots)

It also seems like the key parts of the seed will be within the seed coat; it's not always obvious how to remove this seed coat, or what methods exist. Are there resources for this sort of thing?
As a more concrete example, I've identified this Cardamine hirsuta from my backyard, but I'd like to verify some features of the seeds. Unfortunately, the seeds are very tiny, as this microscope photo of the silique shows, and the features are non-distinct.
Flora of North America provides these details for the Cardamine genus:

Seeds uniseriate, flattened, usually not winged, rarely margined or winged, oblong, ovoid, or globose; seed coat (smooth, minutely reticulate, colliculate, or rugose) mucilaginous or not when wetted; cotyledons accumbent, rarely incumbent

How can I figure out from dissection whether the cotyledons are accumbent  or not? At least, that's the sort of question I'd like to be able to answer.

I will answer my own question for anyone that's interested in doing something similar. First, a great reference for seed morphology is A. C. Martin's The Comparative Internal Morphology of Seeds (1954), available on JSTOR and from sci-hub.
To dissect a seed, you’ll probably want to have a good dissection microscope, two pairs of sharp tweezers, a razor blade, and a pin. Begin by opening the fruit to expose the seeds; for a silique, it’s very easy to peel the valves open. Be careful not to crush any seeds. Using the tweezers (and optionally the razor blade) separate a seed from the fruit by breaking or cutting the funiculus. Do this for at least a few seeds, more is better. The seed coats will protect the seeds from drying out, but be aware, once the seed coat is removed, the embryo will only last a minute or two before becoming completely dried out. You'll figure this out pretty quickly, if it's a problem, otherwise don't worry.

Seeds in the silique, all still in their seed coats.
Use the tweezers to position a seed so you can cut it with the razor blade. To get a good idea of how the embryo is arranged requires several dissections, cutting the seed in different ways (vertically, horizontally, parallel to the work surface); seeds are cheap so just experiment. Cut the seed open and you should have something that looks like the image below.

Seed of Cardamine hirsuta cut in half.
What you’re seeing is (a) the seed coat, and (b) the embryo. Use the tweezers and/or the pin to get the seed coat away from the embryo with as little movement as possible to preserve the formation.

Using a pin, separate the embryo from the seed case. Note that the distinct features of the embryo are becoming visible.

With the seed coat (pale green part) removed, the radicle and the cotyledons become visible and distinct. This seed is accumbent.

The two cotyledons are pressed together in this example, but you can still see a separating line between them when you hold the embryo at the proper angle.
When dissecting seeds, do yourself a favor and select the most mature fruit available. Your life will be much easier if you have a larger specimen. These stills were taken from a video I made, available on YouTube here.

Thanks for the excellent breakdown. IMHO seeds get ignored a lot in identification. This is fair at least where I live in California--the Jepson Manual revolves around reproductive organs. But I constantly encounter fruit & seeds and like to study them as well. What microscope did you use?
Edit: fixed typo