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Different hominid species or simply mutated humans?
I am not a biologist, however, I am curious as to how do evolutionary biologists classify a newly discovered hominid fossil as a separate species. If simply the bone structures are a criteria to classify a hominid specimen as a separate species, how do we know that the different bone structures are not a result of a genetic disorder or a terrible mutation in a human? From what I understand, DNA specimen do not survive that long for consideration. Can anyone provide me with a layman explanation of the process of classification which seemingly doesn't involve genetic data?

As a hint towards the homework aspect - you should look up comparative anatomy and understand what differs in a pathologic (i.e. mutant) vs normal state, and how anatomy might differ between humans and other species.

@bob1 How are we sure that what we consider anatomy of a different species is not a pathology in a human?

Ok then - how do you (as in you personally) tell that someone has dwarfism is human vs non-human? Now, look at the proportions of individual in this state in any animal and extrapolate to other populations and think a little about what we might commonly find as remains?

@bob1 Or rather how do we distinguish between a human toddler and a dwarf of similar height (or built?) based on purely skeletal remains? From what I understand there seems to be some level of guesswork and arbitrariness in how we decide and define a new hominid species which is kind of bothering me.

@bob1 And from what you are suggesting, there seems to be an involvement of extrapolation based on the current diversity of primate anatomy. I was under the impression for a while that there exists theoretical models that allowed biologists to predict the anatomy of our hominid ancestors which are later confirmed or refuted accordingly by discovery of new fossil evidence. But from what I searched on google there is no such methodology under implementation as such. Now I am kind of bothered about this whole thing.

That's where the comparative anatomy comes in - how do pathologists determine the age of bones of a body? (hint: plates) This tells us age. in addition pathologies such as dwarfism have characteristics - look for skeletal pictures for both normal and pseudoachondroplasia or even achondroplasia. This will get you a fair way along to an answer.

The extrapolation was that pathology is rare in populations generally. Bones we find are most likely from "normal" as they are much more abundant...

@bob1 Hmmm. Fair point.

Hey, quite tricky to correlate but good logic
When a fossil is being examined, carbon dating is one of the methods used to examine its age. Also, as time passes, organic components of bones are slowly decomposed and get absorbed in the surrounding (into soil) and the entire mass either collapses under pressure or, inorganic materials from its surroundings replace the empty spaces(rest of the body degrades very quickly). Thus if the fossil is determining exact skeletal structure, then it must have inorganic compounds which resemble to the surroundings. If not(it has inorganic materials that are as same as the inorganic materials of an actual bone), then the fossil is unfit for determining organismal structure, bcuz it had collapsed due to external pressure.
 Now coming to your question.... for mutation to take place, there is a certain time period with a certain percentage of accuracy. Which means that for such a change that causes gross phenotypic difference, it requires a certain number of generations and fixed progeny-coincidences. Thus we can compare the age of fossil and the approximate probability of a mutation, which caused phenotypic changes in a normal human to look like the one determined by the fossil. By this we can calculate the probability that whether the fossil is a legit "non sapien hominid" or just a mutated human.
And often these probabilities are are very low. So we finally identify the fossil to be of a certain hominid with a certain level of similarity to the modern day human.
Hope you understood the process.
References: https://www.sciencedirect.com/topics/biochemistry-genetics-and-molecular-biology/fossil-hominid#:~:text=Most%20early%20fossil%20hominids%20appear,9.10).
https://byjus.com/physics/types-of-fossils/
And for further information on fossilisation and chemical nature of fossils, you can check out this pdf:
https://egyankosh.ac.in/bitstream/123456789/69611/1/Unit-9.pdf

Can you provide any references for this process?

References? Like you want to know in the form of examples or a deeper explanation of the methods and the process?

Yes. Any references or lecture notes perhaps will be helpful.

I've attached the links in another answer...

The oldest DNA from Africa is 15000 years old. From Europe it's 120,000 years, and from Siberia they have horse and mammoth DNA from molars that are [700,000 to 1,2 milliom years old][1]. It's feasible that they will find homo erectus and habilis DNA from cold regions.
The more bone and tooth fragments the better. Then a comparative anatomist can study the age and differences of the bone fragments and articulations to see if they have a lot of similarity with a currently known homo species.
If they can find enough differences in bones of a specimen, concensus can agree that it's a new species. It takes quite a lot of bone to decide that. a small shard isn't enough. Half a skull/leg is fairly vague evidence and half a skeleton can be generally convincing.
[1]: https://www.science.org/content/article/mammoth-molars-yield-oldest-dna-ever-sequenced#:~:text=1.2%2Dmillion%2Dyear%2Dold,record%2C%20reveals%20new%20mammoth%20lineage&text=A%20genetic%20analysis%20of%20long,the%20oldest%20DNA%20yet%20sequenced.


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Can bees find alternate sources of water which mosquitoes can't use?
I live in an area where mosquitoes are a disease vector of concern, and I was dutifully sweeping out a puddle when I noticed a lot of bees had gathered there to collect water.
Is this effort to eliminate mosquitoes also harmful to bees?  Is there water I should leave alone for the benefit of bees without risk of increasing the mosquito population?

Did you do a search using the phrase, "Do bees need water?" This is a site for students, motivated people interested in Biology, and professionals. Oddly for a Q&A site, we require evidence of a search for information in an attempt to answer your own question (a link to a site and a quote is fine.) And fyi, it takes mosquito eggs 6-7 days to emerge from standing water (e.g. puddles) as adults under ideal circumstances. Fresh water in a clean vessel changed daily will not allow mosquitoes to hatch.

Yes, I did.  That didn't tell me much more than I already knew given I had already seen bees collecting it.  Only why they wanted it, and not much on what they need to be able to collect it safely or how that contrasts with mosquito habits and public health information about what we need to get rid of.

If there are running water sources around, the bees are fine. Kinda weird so many bees gathered though so maybe there weren't other sources.

source: Stack Exchange

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

sort: votes


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Found possible bone on St Pete beach in Flordia
What is this bone from? Looks like a vertebrae but not sure. What marine animal could it be from? It’s about an inch Long.

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


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Is the 5% of the oxygen used in the breathing process fully transformed into CO2?
More specific, I found out that the amount of oxygen we inspire is 21% from the total amount, with 0.04% CO2. The expired amount is 16% oxygen, with 4% CO2. Where does the remaining 1% oxygen go? Or are these only approximations?

When you write the number "21%", it represents equally all numbers rounded to 21%; that is, 21% is the same number as 20.5% and 21.49% under the most common rounding convention.
When you subtract two rounded numbers, you can easily get a number that is not zero. For example, 10.500 - 10.499 = 0.001 before you round, but if you round first, 11 - 10 = 1. 10.501 - 10.500 has the same difference, but if you round first and subtract you'll get 0. In other words, 16 + 4 = 21 is completely reasonable math in the world of rounding, the value of "1" is not meaningfully different from zero on that scale.

Got it. So it was just an approximation. Thanks!

@me1234 I wouldn't call it an "approximation" per se; it could be that the numbers were measured very very precisely, but if they've been reported to you as a number like "16%" you should know by looking at that number that it's clearly been rounded to an integer percentage value.


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Is there a convention regarding the use of forward vs reverse DNA strand to represent DNA sequence?
I see in this paper the notation IL-1A C[-889]T which suggests that for this gene, the reference allele is C and the variant is T.
However, when I look up the same gene on dbSNP, it says the alleles are G>A / G>C / G>T.
I noticed for a number of other genes I looked up in the same paper, G becomes C and vice-versa, T becomes A and vice-versa. Is it because of different conventions regarding which DNA strand is used to represent DNA sequence (forward vs reverse)?

"Genetic code" refers to the translation from three-nucleotide codons to amino acids. A change in genetic code would be changing the amino acid that is associated with some triplet.

The first is with respect to the transcriptional unit of the IL-1A gene. For example the "-" indicates that it is upstream of transcription start site.
The second is with respect to the GRCh38 build of the reference human genome, which has a defined orientation.
I don't have access to the paper that you link, but the NCBI annotation for the gene indicates that it is on the reverse strand of the reference genome sequence.
That would seem to indicate that the two annotation methods are concordant, if somewhat confusing. One refers to a position's distance from the start of the gene, one refers to a position's distance from the start of the reference chromosome. Those will necessarily diverge in the reference allele for genes transcribed on the reverse strand.
If you are interested in analyzing the behavior of a specific gene, the transcriptional unit is obviously the relevant level of analysis (for example, predicting translated protein sequence). However, if you are interested in defining global genomic coordinates (for example, mapping sequencing reads), the reference genome become the relevant level of analysis. So each is useful for different purposes.
So, in brief, the answer to your question is yes.

When you say "those will necessarily diverge" do you mean the reference allele will always obey the base pairing rules - like here C for the reference human genome and G when reading from the transcriptional unit ?

@Youcha Assuming that I understand correctly, then yes. C will only ever base pair with G (or at least that's a foundational assumption of how we represent genomes on computers)


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Why are polar seas rich with nutrients and plankton?
It is well known that baleen whales migrate to the polar seas to feed on plankton and krill (marine crustaceans). The question why are the polar seas rich with krill and plankton? If the answer "it is because they are rich with nutrients" then I ask why are they rich with nutrients?

Water density and temperature is very important for understanding water ecosystems. Water is most dense a little above freezing temperature; when surface water is warmer than that, it's less dense and sits on top of cooler water. When surface water is chilled to freezing and increases in density, it can sink and mix with deeper water.
Photosynthesis can only occur in the top of the water column because light doesn't penetrate very deep. When the surface water stays warm and doesn't mix, that top layer gets depleted of nutrients. Mixing brings up anything that has settled and allows life to use the "full column" of nutrients.
See for example https://www.nature.com/scitable/knowledge/library/the-biological-productivity-of-the-ocean-section-70631438/ :

In the high latitude ocean, surface water is cold and therefore the vertical density gradient is weak, which allows for vertical mixing of water to depths much greater than the sunlit "euphotic zone" as a result, the nutrient supply is greater than the phytoplankton can consume, given the available light


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

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

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

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


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How do Addax defend itself from predators?
The addax (Addax nasomaculatus) is a critically endangered antelope adapted to live in extreme desert conditions. It has long impressive horns that can potentially serve as a weapon because they are sharp. Does addax use its horn to fend off predators?
I read somewhere that addax is not a fast animal so trying to escape by running does not seem like a good option for defense.

Picture of Addax and its horns, taken from Wikipedia, photographer: Zachi Evenor
https://commons.wikimedia.org/wiki/File:Addax-Jerusalem-Biblical-Zoo-IZE-611.jpg


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What kind of Bolivian spider is this? Is it a black widow?
I found this spider near La Paz, Bolivia. I have been looking all over the internet to try and identify it, but with no luck. Could someone please help me?
Initially I thought that it was a black widow, due to the red marks and the body shape, but what threw me off is the lack of a distinct hourglass and the yellowish stripes in the middle of the abdomen. Also, the top of its abdomen is mostly red, much more than what is typical for black widows as far as I know. Also, the web doesn't look like a typical black widow web…
The locals told me that they think its a black widow, and that would probably be my best guess, but I'm not 100% convinced… if anyone knows, please let me know!
I have attatched 2 photos below:

Black widows (Latrodectus sp.) are in the Family Theridiidae, or Cobweb Weavers. Your spider's web is an orb web. Cobwebs are messy tangle webs while orb webs are the spiral-like planar polygon figure, sometimes completely vertical and sometimes at an angle, depending on which orb-weaving family made the web. So this spider is clearly not a Black Widow from the web alone. Further, your spider's legs are too short to fit the profile of a Latrodectus.
Searching iNaturalist records for Araneoid spiders (spiders that typically live suspended in webs) in Bolivia, we see the second most-commonly observed spider is one that is black and red.

The second-most observed spider here is from the Alpaida genus, which is in the orb-weaving family Araneidae. And looking at the hundreds of photos available for it, it does indeed build orb webs. However, the ventral markings in the photos of A.versicolor do not match your photos and the legs of A.versicolor are striped/banded while yours are not.
In many spider families, it is difficult to get an identification to species level through photos. But we can try: browsing those top 15 most-observed Araneoid spiders image in Bolivia, we see a number of species in this genus:

Alpaida versicolor
Alpaida veniliae
Alpaida carminea
Alpaida bicornuta
Alpaida rubellula

Some of these look drastically different from yours. But A. carminea matches the general shape, colour and ventral markings.
I am very confident this is from the genus Alpaida, and I lean towards A. carminea but it could still possibly be A. versicolor or some less-common species. A photo of the dorsal (top-side) markings would help distinguish these species.

Thanks! That makes sense yeah, in hindsight I am very upset with myself that I didn’t get a picture of topside markings haha, but the spider’s web was very close to the ground. Anyways, thanks for the help!


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What is a polar plot of relative neural firing
I am taking my PhD qualifying exams on monday, and there is a seemingly simple practice problem that I can't seem to figure out, and I was hoping someone here would have some knowledge, or at least be able to point me in the right direction. My background is in electrical engineering, so apologies for my ignorance.
The question is:
Cell A is strongly phase-locked to spindle oscillations (12 Hz) at 180 degrees, and cell B has monosynaptic excitatory connection to cell A. Assume the synaptic delay is 3ms.
1- Draw the polar plot of cell A’s firing relative to spindle oscillations.
2- What is the relationship of cell B to spindle oscillations? If it is also phase locked to spindle oscillations what would be the degree of locking? Draw the polar plot. If the independently calculated polar plot of cell B does not show significant phase locking with spindle oscillations, what is a possible explanation?
I can't find a single resource that tells me how to draw these polar plots. Can anyone help me out?
EDIT:
12 Hz --> 83.33 ms period
Cell A fires at 83.33 ms * (180deg/360deg) = 41.66 ms
Cell B fires at 41.66 ms +3 ms = 44.66 ms
Phase of Cell B firing is (44.66 ms/83.33 ms) * 360 deg = ~192 deg
Is that right?

Can you explain what you understand and don't understand? Surely you've encountered polar coordinates in engineering, no? have you tried Googling something like "polar plot" + "spindle oscillation"

If you map every spike of a neuron to the phase of an oscillatory process (in your case a 12 Hz sine wave) then you can plot an histogram of that phases. It's usual to plot the histogram in polar coordinates as in the figure below.
For cell B, you will need to map the synaptic time delay in to phase delay using the oscillation period. This is assuming that a synaptic event from cell A induce cell B to spike and there's no other synapses or intrinsic properties of Cell B that perturb the entertainment.

Mansouri, Farrokh, et al. Frontiers in neuroscience 12 (2018): 877.

thank you so much! I couldn't figure out how to add images to this comment, so I added the polar plots to the original post. Does that seem right? I just dont want to misunderstand.
Thanks again


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Is there any objective way to describe ethnicity?
Is there any objective way to describe ethnicity that does not rely on arbitrary sociocultural concepts like referring to racial constructs, modern nations like “Spanish”, or even continents like “European”, but instead just directly describes what traits/genes you have (skin color, facial structure, etc.)?

Please define ethnicity as you understand this relatively recent term, and explain how it differs from the term race, and is not a “socio-cultural concept” too. Examples are really needed if one is to attempt a biological (I assume this is what you mean by “objective”) definition.

Ethnicity is a sociocultural concept. A description of ethnicity without sociocultural concepts wouldn't be objective, it would be wrong.

Ethnicity is a spectrum, with phenotypic stereotypes and with legacy sub-species based on geographical boundaries, which are unacceptable to call subspecies because humans are not animals, sidespecies would work better, although even then... Before 1000 years ago, there were thousands of tribes in relative geographical isolation, even in Europe, and Africa, which didn't have major mixing with other tribes enough to make them similar, for example the Mbuti pygmies of the Congo Rainforest, previous to agriculture booms, almost the entire world was highly differentiated in that sense.

UN 1950 Declaration
As defined by the UN in 1950, 'race' refers to a biological concept while 'ethnicity' refers to a sociological concept. The same declaration said that the way people used the word 'race' was so out of line with biology facts, it would be better to stop using the word entirely:

To most people, a race is any group of people whom they choose to describe as a race. Thus, many national, religious, geographic, linguistic or cultural groups have, in such loose usage, been called “race”, when obviously Americans are not a race, nor are Englishmen, nor Frenchmen, nor any other national group... Because serious errors of this kind are habitually committed when the term "race" is used in popular parlance, it would be better when speaking of human races to drop the term "race" altogether and speak of ethnic groups. [Italics in the original]

Even that document is wrong however, in that it still proposes a definition of race could be rigorously defined, but we now know that is untrue.
Phenotypes
A phenotypic definition of race would determine if people were part of the same race using expressed traits. This basically amounts to phrenology, or the methods of phrenology applied to other traits, most notably skin color and intelligence. Even by 1950 most of these traits were shown to result from environment (e.g. nutrition, education) and not something inherent.
Genealogy
A genealogical definition of race would determine if people were part of the same race using family trees. In 2004 Nature published a study showing that the most recent common ancestor for all living human (the most recent person to shows up in every family tree) was probably from Eastern Asia and lived in ~50 AD. It further showed that the identical ancestors point, beyond which the family tree of everyone alive becomes identical, is ~2,100 BC. As the authors put it, "no matter the languages we speak or the colour of our skin, we share ancestors who planted rice on the banks of the Yangtze, who first domesticated horses on the steppes of the Ukraine, who hunted giant sloths in the forests of North and South America, and who laboured to build the Great Pyramid of Khufu."
Genetics
A genetic definition of race would determine if people were part of the same race using genetic similarities. The problem with this is that Africans are so much more genetically diverse than anyone else. One 2002 study even found the Africans have more in common with Eurasians than with each other. There has actually been a recent push to stop using race as a factor in genetic studies.
How should we understand race
The Stanford Encyclopedia of Philosophy as an article on race, which is a good place to start although it can get a little technical. It also has the section 'Race verses Ethnicity', which you would probably find interesting. Another interesting resource is The Price of Whiteness by Eric Goldstein, which describes how whether Jews were considered part of the White race changed in American multiple times, but always to the benefit of the upper classes.

Race and Ethnicity are not biological traits. They are both inherently sociocultural concepts. They often have biological components associated with them, and sometimes (not always) those biological features help to define a person's race or ethnicity (more commonly with race). But they always have a social/cultural component. And to be clear, if someone is of European or African decent, that is an objective statement about them, even if it's not the most descriptive biologically.
The more objective biological traits can always be studied or talked about in isolation, and then related back to the non-biological components based on their frequency or infrequency in those populations. This is common in public health research. For example, you could talk about specific allelic signatures that occur more commonly in members of the Greek diaspora, or the frequency of sickle cell disease in people whose ancestors came from sub-Saharan Africa.
My primary source is the cultural anthropologist that I'm marrying in a few months. When we met, she taught a university course on the subject of Race, Ethnicity, and Biology, so we had this discussion many times early on. This natgeo article also does a fair job of explaining it in general terms.

How can you answer a question in a scientific manner if you do not define the term that you are discussing? Citing an acquaintance as evidence compounds the unsatisfactory nature of this answer. Other defects are misinterpreting the question as about "biological traits" — a term that the poster does not use. Until the poster gives examples of what he means by ethnicity, it is better to ignore the question.

@David Thanks for your very useful feedback. If you think the question should be ignored due to incompleteness, you should vote to close and explain why in a comment so that others may be convinced to do the same.

I commented to the poster with a request for specific clarification, as is one of the stated purposes of comments. It is only when that is not forthcoming that I vote/ have voted to close, choosing as the reason “lacks detail or clarity” which requires no further comment on my part. As you had answered the question I felt it appropriate to point out that you too were discussing something without defining it, and that I felt your answer unsatisfactory for that reason. Again, I was asking for clarification but from you, and may downvote / may have downvoted on that basis.

Describing ethnicity can be a complex and multifaceted topic. It often involves elements such as cultural practices, language, religion, shared history, ancestry, and sometimes geographical or national origins. While ethnicity can include objective elements like genetic markers or historical data, it's often self-identified or identified by others based on a mix of subjective and objective criteria. There isn't a universally accepted objective method to describe ethnicity because it encompasses a wide range of factors that can vary in significance and interpretation across different contexts and societies.

ethnic classifications based on "objective" genetic data are themselves subjective. there are a lot of subjective decisions that go into how you do that work.

Like the poster, you do not define or give examples of what is meant by ethnicity, so  your post is futile. Please finish the [Tour] and the Help on answering questions. Your answer is just an expression of opinion with no evidence in its support. This is not appropriate for this scientific question and answer site.


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Problem during DNA extraction from blood using automated magnetic beads extraction protocol
We've been using the Perkin Elmer Prepito D instrument + the Blood 250 associated kit for DNA isolation from blood for the last couple of years. It's been working well, however, lately the resulting extracts all show a null concentration. We've tried changing the kit and working with fresh blood with no results. We've also observed that the DNA seems to stay in the disposable deep wells plate bonded to the magnetic beads and it does not get transferred to the final tubes.
Does anyone have experience working with the Prepito instrument or have experienced a similar problem with any magnetic beads DNA isolation protocols ? Does anyone have any suggestion to what might be causing this problem or how we may solve it ?

Have you tried contacting the customer service about the issue?

I agree with @Chris, but there a few things to test first - ensure that the kit is prepared correctly using fresh reagents - don't let a newbie do it!


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Is it possible to use a fluorometer such as Qubit or Denovix DS-11 FX to detect DNA integrity/fragmentation?
I remember someone saying that the Thermo Fisher Qubit can be used to assess DNA integrity, is this true?
Our lab recently acquired a different fluorometer, the Denovix DS-11 FX. I'm wondering if it can be used to detect DNA integrity. I couldn't find any info about it in the manual.

In general, methods like Qubit rely on specific interactions between a fluorescent dye and the target molecules, be they dsDNA, ssDNA, RNA, or whatever. These are sensitive to the overall amounts of targets, not their size distributions (within reason, of course). For DNA, you will need to use an electrophoretic method like a Bioanalyzer or an agarose gel to quantify integrity.

The Qubit has an RNA integrity kit. They call it RNA IQ which works off binding of two different molecules. I haven't used it, so can't say if it is comparable to bioanalyzer, but they claim it is as good as electrophoresis (presumably visual inspection for integrity on this one). Bioanalyzer is the gold standard for RNA AFAIK, don't know about DNA.

There isn't an equivalent fluorescent method for DNA. Bioanalyzer orTapestation is the way to go.


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Speed of cell signal to protein production
How long does a cell take to make a protein from the point of receiving the signal to produce the protein? Does the cell have stores of the protein ready to release or is the protein created in real-time (transcription -> translation -> protein release)? Does this vary depending on the type of protein?
I have read articles that describe transcription as fast as 40 - 80 nucleotides per second. The same articles described translation as taking a similar amount of time. Taking into account the time it takes to move the mRNA from the nucleus to the ER and then to the point where the resultant protein is ready to secrete, how long does it take?

Totally depends on the pathway and protein and what you measure.

@BryanKrause What are some general estimates? Perhaps an example of a specific protein?

Milliseconds to days.


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Are there any known bioluminescent species of fungi found in the Pacific Northwest?
This might not be the best place to ask this but since mycology is included I though I'd try: Some 21 years ago in late May of 2003 I was walking with a group of friends in the woods near Olympia, Washington sometime after 9PM when I noticed a blotch on the trunk of a tree(the tree was standing) glowing blue-green. It's glow was much dimmer than a firefly but it was clearly visible. We had no flashlights or lighters and it was pretty dark so I couldn't see what it actually looked like during daylight. It appeared to be either a slime mold or a crust fungus but I didn't touch it so I have no idea what its texture was. I talked to someone who also saw something very similar in the woods near Astoria, Oregon. So are there any known slime mold or crust fungus species that are bioluminescent? Nobody seems to have any idea what is could be and it makes me wonder if it is some species that has yet to be officially calatoged by science.

https://www.reddit.com/r/mycology/comments/5o1mry/do_any_bioluminescent_mushrooms_grow_in_washington/

@BenBolker it clearly wasn't any of those. I would say it was not a mushroom in any sense of the word but more like a mold of sorts if you will. If I ever see it again, I will take a sample of it to a mycology lab.


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Can anybody identify what insect laid these eggs? Find in my backyard in N. Texas
I happened to turn over this leaf in my backyard and found these eggs.  Would really like to know what they're from. I'm in North Texas.

Maybe https://springfieldmn.blogspot.com/2018/11/fuzz-on-kale.html


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Has artificial selection / selective breeding ever been targeted at intelligence?
Has it ever been attempted to breed a species, for example dogs, to maximize some aspect of intelligence? I find it particularly interesting to imagine one could selectively breed dogs to be able to talk or develop far more advanced communication structures.
Is there anything we know that could help us predict how many generations of breeding this might take?
Has this idea even been abstracted or generalized to a general technique of “targeted breeding”, where, in a highly methodical and systematic, scientific way, someone pinpoints some specific, even measurable trait, and then with as much automation as possible, tries to artificially breed many generations of an organism to continually maximize that trait, as fast as possible?

when you researced this what did you find?

I'll briefly note that when applied to a human context (which is the most common for this question), this kind of breeding is called "eugenics" and usually frowned upon. Assuming that you are interested in breeding dogs, I'll point you in the direction of the breeder's equation. But I'll also point you to the wiki page on IQ heritability (in humans), which has a number of important caveats. IDK if they apply equally to dogs.

Hi! Many dogs have been bred to be more alert and bark a lot rather than be bulkier and more aggressive, precisely using a systematic breeding method, maybe not as precise as scientific research. Would that count? This question is hard to answer without a common reference framework for intelligence.

@MaximilianPress - It must pertain to dogs as well. I bred Border Collies, and while I didn't employ a scientific method, some were more "intelligent" than others, and one of them was absolutely outstanding, demonstrating easy familiarity with syntax and concepts.

your last paragraph more or less exactly describes the process of artificial selection, which is being done all the time in agricultural - selecting animals or plants for increased crop yields, etc etc.

Has it ever been attempted to breed a species, for example dogs, to maximize some aspect of intelligence?

Yes, perhaps unintentionally, but certainly to serve a purpose: to better herd sheep.
Border Collies (BC) are widely considered to be among the most intelligent of dog breeds. For (at least) hundreds of years, they have been selectively bred to herd sheep, playing on a dog's natural prey drive.
By listening for and obeying different whistles/verbal commands for different actions, a working BC at minimum learns whether or not to approach the flock, find the sheep, gather them into a group, move them towards the shepherd or not, towards the right, towards the left, leave the flock to go back for strays, speed up, slow down, walk, stop, lie down, bark, and come back (you're done.) There are also other commands, but those few are the minimum. (I've seen a BC cut 3 sheep by name out of a flock of ?50-60 sheep and herd them into a separate enclosure. They were not marked.)
By selecting, for generations, the most responsive dogs to breed, BCs have become finely attuned to look to humans for commands of all kinds, and to obey them.
While they may not ever learn to talk (not only vocal cords, but the entire mouth including most of the tongue is needed for speech), they can still communicate. A BC named Chaser was trained to learn the names of over a thousand toys, many of them being similar (e.g. red ball, fuzzy ball, big elephant, little elephant, white elephant, etc.). The trainer, a psychologist, stopped at 1022 objects because of space and he was kind of bored at that point. The dog could find a photograph of the object by name, and could find on the first try without help an unknown toy in a group of familiar toys when the owner made up a new name, understanding the concept of unfamiliar. He taught her verbs, adverbs, adjectives, and syntax. She could follow a string of commands successfully.
Word buttons are now being used to study canine cognition, the most well known example of these dogs being Bunny the Sheepadoodle. This video explains Bunny's progress and ability to communicate using abstract ideas, like "ouch" and "stranger", and emotions. This is as close to talking as we are likely to get with dogs.
Sadly, Bunny often indicated that she was sad, very sad ("Big Sad") and worried ("Concerned") and has asked for medications for relief. Scout's honor. (How 'human' is that?) Personally, I can't understand someone doubting the capacity for intelligence (and emotions) in dogs after seeing these dogs in action.
If you can stand another anecdote, read on. Border Collies are "soft dogs", meaning they don't like hard corrections, e.g. they react negatively at being yelled at. My grandfather, a farmer who had a small herd of dairy cows, could tell his Border Collie which cows he wanted her to round up and bring in at milking time, and she would do so. In the summer, cows are milked twice a day. One summer evening she missed a cow, which is kind of a big deal, as missing even one milking can decrease milk production for days. Uncharacteristically annoyed, my grandfather angrily yelled at her to go back out and get said cow, which she did. She then disappeared for 24 hours, making my grandfather go fetch his own damned cows from the fields. Did she disappear because her feelings were hurt, or to teach him a lesson? I don't know, but my grandfather never yelled at her again.

The Wikipedia article on "Selective breeding", which I went to while composing a comment on your question, has this sentence:

A Soviet attempt to breed lab rats with higher intelligence led to cases of neurosis severe enough to make the animals incapable of any problem solving unless drugs like phenazepam were used.[14]

The reference given is in Russian (see here), but even with Google Translate it's hard to get much out of it.
A Google search on "'artificial selection' learning evolution" brings up a lot, but nothing that I can find specifically on selection for intelligence: there are experiments on artificial selection for brain size in guppies, reversal learning (cognitive flexibility) in birds (Parus major), learning rates in parasitic wasps ...


text:
How quickly new infectious viruses can be identified and isolated today?
Say, tomorrow, a previously unknown disease (with non-unique symptoms) starts spreading: With the techniques we have today, how quickly can we find out which virus might be the root cause?
I am asking this, as I was reading this article about the discovery of HIV:
https://www.pasteur.fr/en/research-journal/news/40-years-hiv-discovery-virus-responsible-aids-identified-may-20-1983
Apparently, it took several years after the epidemiologic observation of AIDS till HIV is isolated and identified as the root-cause. In the case of SARS (2002), this process took less than 5-months:
https://www.ncbi.nlm.nih.gov/pmc/articles/PMC7112372/
And more recently, for SARS-CoV2, (apparently) it took only 12 days from the first announcement to the genetic sequencing of the virus:
https://www.who.int/news/item/27-04-2020-who-timeline---covid-19
Compared to months/years, identification of the COVID 19 virus within days looks impressive: In the future, can we expect to routinely identify any new infectious virus within days? or cost/effort for these techniques still limit their wider and routine usage?

Here's a hit: technology, specifically sequencing.


text:
How to understanding the mouse strains name and differences in mouse strains used in biology
I am currently working with various mouse strains in my biological research and I am encountering some confusion regarding their nomenclature and differences. I understand that the different names indicate different origins, but I am finding it difficult to decipher the specific meanings behind these names.
For instance, I frequently come across the strain C57BL/6JN. It seems that C57BL represents a common mouse strain, but I am unsure about the meaning of the slash and the 6JN. Could someone kindly elaborate on this?
Additionally, I have seen the name C57BL/6NJ referenced in some literature. Is this the same strain as C57BL/6JN, or do they possess certain distinct characteristics? I would appreciate any clarification or guidance on this matter. Thank you!
Regrads

There once was a mouse numbered "57". Someone used that mouse to make a bunch of inbred lines. One of those lines, number 6, survived well. It had a black coat.
This is the C57, black, line number 6, or C57BL/6, also called "black 6". Remember that inbreeding is typically "bad", so it takes a lot of tries to make an inbred mouse line; most turn out to have fatal diseases or poor reproduction.
The original line was and is maintained by Jackson Labs, we call that strain J. NIH started maintaining a line from that same stock. We call that strain N or 6N; sometimes JN because it started as J. These are all inbred lines so they "should" be identical, but in practice mutations happen and genetic drift can occur; this is mitigated by back-crossing within the inbred line but Jackson wasn't back crossing theirs with NIH and vice versa, so it's proper to consider them separate lines.
Jackson Labs also now has taken the NIH strain and maintained it to sell:
https://www.jax.org/strain/005304
They call this C57BL/6NJ: the 6N strain now back at J.
Other people have taken and bred lines from these strains so there are numerous others, most are less significant since they're not mass produced commercially. Jackson Labs has really good documentation on their mouse lines including known SNPs between lines, if you're ever curious I'd start there and see what they have documented before looking for other sources.
Some other references:
https://en.m.wikipedia.org/wiki/C57BL/6
https://www.jax.org/strain/000664
Bryant, C. D. (2011). The blessings and curses of C57BL/6 substrains in mouse genetic studies. Annals of the New York Academy of Sciences, 1245, 31.

Thank you for your answer that's awesome! Can I ask an additional question to confirm my understanding? Taking C57BL/6NJ and C57BL/6JN as examples, first, Jackson Labs obtained a well-characterized mouse strain C57BL/6J through inbreeding up to the sixth generation. Then, NIH acquired and maintained this strain, so the strain maintained by NIH is referred to as C57BL/6N or C57BL/6JN. C57BL/6N and C57BL/6J have the same genetic background (except for genetic drift and new mutations that may occur after separation).

Afterwards, Jackson Labs obtained C57BL/6N (also known as C57BL/6JN) from NIH and commercially sells these mice, which are referred to as C57BL/6NJ

@zhang Nothing about "6 generations". Dozens of generations. 6 is not the generation number, it's one of several lines started from the same mouse.

@zhang inbreeding typically goes on for 20+ generations to fix the genetic lines. The 6 refers to the original line - it'll be 6th attempt or 6th mouse pairing they tried to do this to, or something similar. Nothing to do with generations.


text:
What can't the human eye see far-UV, X-rays, and gamma rays?
This is as much of a physics question as it is a biochemistry question. But how does rhodopsin(and the retinal chromophore itself) respond to ionizing radiation? Photons whose wavelength is longer than 700nm do not have the energy to photoisomerize retinal. But for photos with wavelength shorter than 300nm for sure have more than enough energy to do so.


text:
Can contaminants in the DNA extract disturb the sequencing?
I'm having troubles with some DNA extractions. The yields are much lower than expected, and the Nanodrop spectrophotometer is showing quite a bit of contamination through the 260/230 ratios (see table below). The samples consist of mixed marine fine-mortarted fauna (environmental samples), of which I want to sequence a couple of marker genes to determine species composition (metabarcoding), using paired-end Illumina MiSeq. I could just go on with the PCR as I'm supposed to and see if that works, but my concerns are still:

As the DNA extracts contain an order of magnitude less DNA than expected, would you really consider them representations of their whole communities?
Assuming the PCRs work out and I get nice bands, is there still a risk that the contaminants could disturb the sequencing process? That is, given that I only input 2 out of 25 µl of DNA extract into the first PCR reaction, and just as little PCR product into the second (index/library prep) PCR. Can the Illumina flow cells be sensitive to these contaminants as well?

Sample ID
Nucleic Acid Conc.
Unit
A260
A280
260/280
260/230
Sample Type
Factor

1
7.6
ng/µl
0.152
0.086
1.76
0.71
DNA
50

2
24
ng/µl
0.479
0.249
1.93
1.16
DNA
50

3
21.9
ng/µl
0.439
0.238
1.84
1.15
DNA
50

4
18.6
ng/µl
0.371
0.190
1.95
1.29
DNA
50

5
18.5
ng/µl
0.371
0.200
1.85
1.44
DNA
50

6
6.9
ng/µl
0.138
0.069
2.00
0.70
DNA
50

7
7.9
ng/µl
0.157
0.092
1.72
0.78
DNA
50

8
35.2
ng/µl
0.703
0.363
1.94
1.67
DNA
50

9
9.6
ng/µl
0.192
0.099
1.94
0.85
DNA
50

extraction_blank
1.6
ng/µl
0.031
0.006
4.84
0.19
DNA
50

collection_blank
3.9
ng/µl
0.078
0.036
2.19
0.42
DNA
50

According to ThermoFisher "good" values are between 1.8-2.0 for the 260/280 ratio, and 2.0-2.2 for the 260/230 ratio. Less may indicate the presence of salts or EDTA for example. The concentrations are for 100 µl of DNA extract, that is, total DNA yield is given by multiplying the concentrations with 100. For the record, I have heard from people who get 200-300 ng/µl DNA concentrations from marine sediment extractions. My samples should in theory contain even more than that. Although I have earlier only gotten concentrations in the range of 30-100 ng/µl for the same type of samples with the same extraction kit.
What would you do in this situation? How do you go on with values like this from the spectrophotometer?

I have never performed environmental extractions, and I am not familiar with your specific protocol. I would defer to any other commenters with more experience with these methods. However, I would recommend redoing the extractions, if it is feasible. Any time a standard protocol gives results that are significantly unexpected, it's worth thinking hard about. In this case, where it is your initial step, it's even easier to suggest, since there's much less wasted time than if it were later in your protocol. The quality of your data depends on what goes in, so it's not worth risk it.
Nanodrop quantification is not particularly reliable for low concentrations like this. That makes the ratios even more questionable. I would not try to make too much of these particular metrics beyond "this was a poor extraction."
PCR could certainly give you plenty of DNA even from a minuscule sample - that's the whole point of exponential amplification! But if your extractions are poor enough that they are extremely biased, that would carry on through PCR.
I would worry less about contaminating the sequencing reaction at the end, but it's still worth trying to get pure DNA. What's your cleanup before dilution? Anyway, if you do have an inhibitor, I would expect it to give issues during PCR.  And you'll be diluting them 20? 100?-fold for loading onto the flow cell.

Hi. I forgot to add that I've redone the extraction four times already, following the kit manufacturer's troubleshooting guide for low DNA yields: heating up the elution buffer, letting it incubate for longer, running the elute through the spin column multiple times. I've even bought a new kit to try with. The values above are as good as it gets. Thought I was gonna do another thread about that... And I've double-checked the concentrations with Qubit, although not for this last specific extraction attempt. It doesn't differ that much.

The point of this thread was more to check which possible unforeseen problems I might encounter later, should I choose to continue with these DNA extracts. I perform post-PCR cleanup using magnetic AMPure XP beads. The final dilution of the DNA extract in the library will probably be closer to 1:100. But I take your answer then that I should not worry about contaminants as long as the PCR bands look nice – I should worry about bias from the low DNA yields?

With a bead clean and that dilution, I wouldn't worry about sequencing. I would still be a bit concerned  about bias. I don't know how quantitative you need this assay to be (do you want to measure frequency or just presence?), but I would be concerned by the extraction issues.

In metabarcoding studies, quantitative (read count) measurements are largely useless due to the large bias introduced by PCR. Presence/absence is the primary goal. I made another thread about the low yield issue: https://biology.stackexchange.com/questions/113837/why-are-my-dna-extractions-failing – Kudos to whoever can solve that mystery.

You might be able to get away with using these, then, but I would definitely qubit instead of nanodrop and be skeptical about any results you get.


text:
Reference request for the evolution of faces
We humans have faces, as well as many other animals, like cats, dogs, monkeys, etc. But there was a time on planet Earth before there were any faces in living things. I am currently very interested in how faces evolved. Is there any book or paper on this topic? I would like some references.

What is and isn't a face? Do mice not have faces? What about a crocodile? A fish? A fruit fly? If any of those do not have a face, what makes what they do have not a face?

To expand on Bryan's question: how is it different from the evolution of heads?

@MaximilianPress A head of lettuce, for example.

If what you are interested in is really the evolution of heads (which I think imply faces), then cephalization is what you are looking for.
Namely, when bilaterally symmetric organisms colocalize sensory organs, feeding apparatus, nervous system, and in most cases a brain.

Fig 1: Wikipedia's figure showing where cephalization occurred.
Note that starfish and other echinoderms lost their ancestral cephalized state and went back to radially symmetric headless animals!

Interesting article in Nature: https://www.nature.com/articles/s41586-023-06669-2 "This finding suggests that from the perspective of ectoderm patterning, echinoderms are mostly head-like animals and provides a developmental rationale for the re-evaluation of the events that led to the evolution of the derived adult body plan of echinoderms." - perhaps they're more bodyless than headless!

@BryanKrause woah. biology is weird!

@MaximilianPress Perhaps you are the one who is weird, he who has a strange body hanging off its head.


text:
How to calculate the osmolality of Hepes solution?
Here is an excerpt from Kollmann et al. (2020, J. Physiol.):

This ring was placed in a recording chamber continuously perfused with 37°C aerated Hepes solution containing (in mM) 136 NaCl, 10 glucose, 5 KCl, 10 Hepes, 1.2 MgCl2, 2.5 CaCl2 (pH 7.40) at a rate of 11 ml min-1. Hypoosmolality of the Hepes solution was established by reduction of the NaCl content to 33 mM (94 mOsm kg-1 H2O), 58 mM (144 mOsm kg-1 H2O) or 83 mM (194 mOsm kg-1 H2O).

For validation, I tried to calculate the osmolality by myself. For example, when the NaCl content is changed to 33 mM, the osmolality should be
$$
33\times2~(\mathrm{NaCl})+10\times1~(\mathrm{Glucose})+5\times2~(\mathrm{KCl})+10\times1~(\mathrm{Hepes})+1.2\times3~(\mathrm{MgCl_2})+2.5\times3(\mathrm{CaCl_2})
$$
which gives 107.1 mOsm/L, i.e., 107.1 mOsm/kg. But 107.1 is largely different from what the author stated, namely 94 mOsm/kg.
When the NaCl content is changed to 58 mM or 83 mM, the osmolality should increase by (58-33)*2=50 mM, or (83-33)*2=100 mM. This is consistent with the author's calculation, i.e., 144-94=50 mOsm/kg, or 194-94=100 mOsm/kg. So, my calculation is correct at least in terms of NaCl. But how can I calculate the contributions of the remaining solutes correctly?
Reference: Kollmann, P. et al. Submucosal enteric neurons of the cavine distal colon are sensitive to hypoosmolar stimuli. J. Physiol. 598, 5317–5332 (2020).

I assume this is a measured osmolality rather than a calculated one. It's challenging to calculate actual osmolality because of incomplete dissociation of the solutes, see for example https://en.wikipedia.org/wiki/Osmotic_coefficient. There's also a subtle difference between osmolality (particles per mass) and osmolarity (particles per volume).
I believe HEPES can contribute to this effect because it will bond with free ions. Sugars do, too. The gap still seems a bit larger than I'd expect, some of that might involve the temperature of water when originally mixed versus when measured. If you're trying to replicate their methods, I'd get yourself an osmometer.


text:
Bacterial cells disappearing from culture with time
While growing my bacterial culture in liquid media for protein expression, I have been observing that the bacterial cells (E. coli) grow well for some till OD600 reaches ~0.4-0.6, then suddenly disappears. The turbid culture having grown cells turned clear, and I could not see significant cell debris precipitated. But I do see some particles that make the media look hazy. Seems like the cells are getting lysed. I have tried changing the media components, resources, temperature, cell lines, genes of interest, etc. BUt nothing seems to work. The pH of the media seems to be the same before and after inoculation and lysis as well. The behavior is quite random and doesn't follow any pattern. Any suggestions on possible causes or similar experiences would be really helpful. Thanks.

That sounds like phage. The pattern of initial growth followed by catastrophic death with some cell debris is classic for induction and lysis.
Have you observed anything strange on plates? You could just do a simple plaque assay using some of your liquid culture to confirm.

I thought of that possibility as well. The transformed colonies on plates with antibiotic looks normal. A few times I have observed wierd colony morphology (not quite circular, but distorted). I did try plaque assay, but couldn't notice any area of clearance on LB-agar plate with bacterial lawn. We fumigated (formaldehyde-permanganate) the lab and all equipment, washed all glassware with bleach, and alcohol followed by autoclaving before preparing fresh media. Still the problem persists. Do you know of any other method to detect/ get rid of phage contamination?

That's tough. I was in a lab that struggled with recurrent phage contamination before, and there isn't a great solution. Fumigation sounds like a good idea - we weren't able to do that. We found a few things seemed to work well enough to keep things under control: this might not work for your combination of phage and cells.
-Virkon is better than bleach. Wash all flasks and regularly wipe down counters,  pipettes, and common instruments and spaces.
-High temperature is better than autoclaving. See if there's an oven you can use to sterilize flasks.

(cont)
-Filter pipet tips may prevent transferring aerosolized phage.
-Try to prevent aerosols in general. Be gentle with your cells and don't leave plates open.
-You mentioned trying alternative strains, but you might look into this a bit more. Some common phage receptors differ between B and K strains, for instance.
-You could try low temperature or cold inductions to potentially reduce induction.
-Regular maintenance: once phage has established itself, you probably want to have regularly-scheduled deep cleaning (especially surfaces where it can linger) to keep it down.

(end)
-Vigilance: you will never remove all the phage, you'll just keep them under control. Make sure everyone watches for signs of lysis and communicates them.

Is it possible that your E. coli strain is a lysogen? Prophage induction at high cell density could explain why your culture is crashing

This might be an unrelated question, but may I know what protein you are expressing ?


text:
Why are my DNA extractions failing?
Alright, here's a tricky case. I am trying to extract DNA from mixed marine faunal samples, something that has been done in my group for a long time, and which I have done successfully before as well. Now it's just not working anymore for my last batch of samples, even though they should be very similar to my previous samples, and I'm using the exact same extraction kit (ZymoBiomics DNA Miniprep kit D4300). The yields are much lower than expected, and the Nanodrop spectrophotometer is showing quite a bit of contamination through the 260/230 ratios (see table below). My goal is to do metabarcoding of a couple of marker genes to determine species composition, so I need the DNA extracts to represent their whole communities. When yields are so low, I believe the chance of severe bias increases. I have heard of people getting 200-300 ng/µl from marine sediment, and my samples should in theory contain even more. Nevertheless, what I have gotten used to from earlier extractions are yields around 30-100 ng/µl (all values based on 100 µl). This is something different though. The table below shows the best results out of four extraction attempts. I have double-checked the concentrations with Qubit and they are approximately the same. I should also note that I ordered a totally fresh kit for this extraction, to rule out that something had gone bad in the previous one. But it's not much of a difference. I have also followed Zymo's advice to increase DNA yield: heat up the elution water, let it incubate for longer, run it through the spin column multiple times. It does not significantly affect the results. We preserve our samples in DESS (20% DMSO, 0.25M EDTA, NaCl saturated solution). Zymo suggested that the high salt content could disturb the extraction process. I'm not too sure, since it has worked earlier. But maybe that could be the reason why we are never seeing >200 ng/µl yields? What do you think?
What makes me believe that it's not the samples that have gone bad, is: a) That they would need to get really damn bad before yielding this little DNA, and b) That between every extraction attempt, there is usually one or two samples that spike and show quite ok DNA concentrations (30-60 ng/µl) – but rarely the same samples.

Sample ID
Nucleic Acid Conc.
Unit
A260
A280
260/280
260/230
Sample Type
Factor

1
7.6
ng/µl
0.152
0.086
1.76
0.71
DNA
50

2
24
ng/µl
0.479
0.249
1.93
1.16
DNA
50

3
21.9
ng/µl
0.439
0.238
1.84
1.15
DNA
50

4
18.6
ng/µl
0.371
0.190
1.95
1.29
DNA
50

5
18.5
ng/µl
0.371
0.200
1.85
1.44
DNA
50

6
6.9
ng/µl
0.138
0.069
2.00
0.70
DNA
50

7
7.9
ng/µl
0.157
0.092
1.72
0.78
DNA
50

8
35.2
ng/µl
0.703
0.363
1.94
1.67
DNA
50

9
9.6
ng/µl
0.192
0.099
1.94
0.85
DNA
50

extraction_blank
1.6
ng/µl
0.031
0.006
4.84
0.19
DNA
50

collection_blank
3.9
ng/µl
0.078
0.036
2.19
0.42
DNA
50

I don't see that this question is significantly different to your previous question, which is essentially the same the question as here. Either way, you will need a bit more detail on the samples - how are they stored, how much input, what is in the samples (tissue? soil?), what has changed between previous extractions and now, have you gone back to old (known working) samples and tried re-extracting?

Does this answer your question? Can contaminants in the DNA extract disturb the sequencing?

I'd suggest trying an alternate sample prep method that is a little easier to introspect than the kit and is optimized for difficult samples. For example this chapter describes a "difficult sample" extraction method that works on soil or fungi, which are notoriously hard to get DNA out of.

Bob: The samples are mixed marine faunal samples. That is, a mixture of anything larger than 40 µm that lives on a settlement plate in the sea, or swims near it at the time of retrieval. Directly upon retrieval, samples are mortarted thoroughly and preserved in DESS as described above, then stored in -20℃. What changed between the previous extractions and the last one is that I got a fresh ZymoBiomics DNA Miniprep kit for the last extraction. For the previous extractions I played around a bit with the tips that I got from Zymo, but the point is that it didn't make any significant difference.

Today I also tried with the Qiagen DNeasy PowerSoil Pro kit, but the results were practically the same, or possibly even slightly worse. So I will go back to the Zymo kit. Maybe redo some old samples that I know have been working before...


text:
Identify caterpillar
Hi i live in the Nilgiris region of South India and I found this fellow on one of my Lemon Grass plants. About 8-10 cm long with fine hair all over.
Let me know what species this could be. Thanks


text:
Why UniRef representative sequences in UniProt are so long?

I found that some sequences in UniRef90 database are tens of thousands amino acids long. I read UniProt official documentation and figured out that these sequences are the representative sequences of clustered known protein sequences. That is, they are sequences of real proteins that do exist in nature.
I know that some giant proteins like Titin have the length of tens of thousands a.a. However, when I visualized the lengths of protein sequences in uniref90.fasta, I got the plot below.

It seems that at least thousand of proteins are longer that 10,000 a.a. and average protein sequence length is around 5,000. So far, I have thought that proteins are hundreds a.a. long. Do natural proteins are usually composed of thousands of a.a?
Thank you.

This is a good question, but there are a few things going on. There is a subtlety about what you're asking: do you want to know the average length of a protein sampled from a cell, or the average length of a sequence sampled from this specific database? Because the plot you use relates to the second question, while you seem to want to know the first one.
The whole Uniref is also probably not the best set of sequences to test this with. The clustering would tend to amplify the proportion of outliers caused by missing stops, skipped exons, pseudogenes, or duplications, since they will form their own cluster. Also, each isoform can form its own cluster. From a quick look at the first ~100 Uniref90 sequences, maybe 80 of them are annotated as titin isoforms. Plotting and comparing the distributions of Uniref50, 90, and 100 might be a good exercise. At the very least, median is better than mean here.
This is a good relevant paper: Protein length distribution is remarkably uniform across the tree of life
Also, this from the always-excellent bionumbers is good: https://book.bionumbers.org/how-big-is-the-average-protein/

Now I understand. The plot I made shows only the representative sequences of UniRef90 clusters. It doesn't consider about the other members in each cluster. Therefore the average length must differ from that of proteins sampled from a cell, which considers any sequences whether it is representative sequence or not. Also, such representative sequences contain may outliers due to the reasons you've mentioned. I will analyze the data again keeping your advice in mind. Thank you so much.

There's something odd about your numbers:
I took a quick look at the database and just used the (LENGTH:[X TO Y]) command to filter by lengths. This gives you numbers of entries for each size bracket (see numbers on left of image under the Clusters heading).

These are the numbers for clusters I get in the UniRef90 database (Yes, there's some overlap in the brackets 'cause I used 0-1000, 1000-2000 etc, but it doesn't matter in the final analysis, all it did was save some typing time for me):

Size bracket
Number of clusters

0 - 1K
173,209,693

1K - 2K
5,303,314

2K - 3K
658,504

3K - 4K
161,999

4K - 5K
80,517

5K - 6K
32,742

6K - 7K
12,207

7K - 8K
7,076

8K - 9K
4,995

9K - 10K
2,616

10K - 11K
1,603

11K - 12K
900

12K - 13K
792

13K - 14K
476

14K - 15K
519

15K - 50K
3902

From this you can see that the first two brackets (even the first one alone) are vastly more than all the others summed together, which means that there's no way that the average cluster can be 5000 aa in length
This tells me that you have some real problems in your analysis or in your data, but what that problem might be, I don't know. I thought it might have been that you had substituted size for length, but even there cluster size of 5K-6K is only in the 400 range and vastly outweighed by the 1-1000 clusters.
Rest assured, the average protein size is between 50 and 2000 amino acids in length (see 4th para in link).
Edited to add:
I did some analysis of my own using R and ggplot. I downloaded a dataset specifying 90% identity and lengths to be between 2000 and 8000, to keep the file smallish. This makes it a fairly small dataset of 952,569 entries and using the following code (yes, poor coding style, but it's literally a 4 line code including loading the file and library (left out here), so don't need anything fancy). The binwidth of 200 means that the counts for each category are "binned" in lengths of 200 aa, meaning there are 5 bars per 1000 aa.
plot <- ggplot(file, mapping = aes(x = Length))
plot + geom_histogram(binwidth = 200)

This shows me that there is a peak number of entries between 2000 and 3000, not the ~5000-6000 peak in yours (I note that your bins don't line up with nice groupings, so it makes it hard to work out what they are, but something like 910 aa per bin I think, so it should be about 5400 at the peak). This peak has about 200,000 entries, which is massive compared to the ~45,000 peak in your plot, and if you pool my numbers into brackets of ~1000, you get an even more telling picture:

These plots, together with the numbers in my table above, tells me that there is a problem with your dataset or the code you used to generate the plot, probably the data table.  You might wish to explore your data more - look at the number of entries in the table. I think, from your plot, that you have vastly too few entries to be a full UniRef90 dataset, as they should total somewhere close to 180 million not the (very rough) <200,000 entries you have.

Thank you for your answer. I downloaded UniRef90 dataset again from the official UniProt website but got the same result. I think there must be some errors in my own Python script which I used to analyze the data. So currently I'm trying to debug it. (It might take some time since I have bad coding skills.) And thanks to you, I became aware of the existence of Uniprot query syntax and was able to learn it. Thank you again, I’m really grateful for your help.

@dearname You are welcome. If you think either of the answers is the correct one, then you should hit the checkmark below the voting arrows to tell people that it is the accepted answer. Stick at the programming, it's a steep learning curve, but is a very rewarding and looks great on your CV. Data science is hard, but it always pays to do some exploratory things (like looking at number of entries, quick distribution plots etc) before jumping in to a full analysis.


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What is meant by the stabilization of a receptor?
I am reading a journal paper, and have a question about the below statement:

PSD-95 is involved in the recruitment, trafficking and stabilization
of N-methyl-D-aspartic acid receptors (NMDARs) and
α-amino-3-hydroxy-5-methyl-4-isox-azoleproprionic acid receptors
(AMPARs) to the postsynaptic membrane during cortical development

What is meant by the stabilization of a receptor? Does stabilization refer to a receptor being protected from being degraded (e.g. lysosomal degradation)? Any advice is appreciated.

The key phrase is "to the postsynaptic membrane" which in that sentence modifies all of recruitment, trafficking, and stabilization.
Recruitment to the postsynaptic membrane means greater concentration of the receptor.
Trafficking to the postsynaptic membrane means externalization from vesicles to the actual external membrane.
Stabilization to the postsynaptic membrane means the receptors stay there and don't get re-internalized.
Though these may seem like they're all the same thing, the responsible machinery is different and the different processes can be measured separately. In the "leaky bucket" model it's the water level, water dripping in from a faucet, and water leaking out from a hole. To be clear, it's unlikely that PSD95 is actually doing all these entirely by itself, rather it's involved in signalling that has all these effects one way or another.


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Overly stabilised population dynamics model
My project aims to model population dynamics of a 3 trophic levels system: primary producers, herbivores and predator. I try to predict how changes in the vegetation abundance might affect higher trophic levels. My herbivores are described through juvenile and adults compartments, with the first "supplying" the other, after a maturation time. My predator only has an "adult" life stage.
Currently, I'm a bit struggling, because no matter what vegetation biomass or initial densities I put in the system, it always converges toward the same densities. The system reaches an equilibrium stable point, no matter the amount of food available for my herbivores, which is quite a problem for my study.
Could anyone enlighten me with some ideas of what to test/look for to resolve my issue?
Of course, I can give more details about my simulations (done with R and Rstudio, with the deSolve package) and the structure of my equations (Lotka-Volterra).
Thanks a lot in advance.

If you're still interested in the answer, can you edit your questions to include the actual equations (either a R code or as mathematical notation, ideally in LaTeX but text would be OK -- just not an image/screenshot in either case) you're working with?

Hi @BenBolker, Yes I'm still interested! I'll try to edit my post as soon as possible. Thanks a lot

In a deterministic system, there are 2 options: equilibrium or collapse. It sounds like you've found an equilibrium. This is probably expected!
Classic Lotka-Volterra makes specific deterministic claims about the equilibrium population levels:

Secondly, the population equilibrium of this model has the property that the prey equilibrium density (given by $x=\gamma /\delta$) depends on the predator's parameters, and the predator equilibrium density (given by $y=\alpha /\beta$) on the prey's parameters.

As long as your inputs are consistent with the model (e.g. nonzero), they should come to equilibrium. You are basically finding the abundance states where change in frequency over time is zero.
I don't know much about the deSolve implementation, but I'd suggest varying not the initial densities but the other parameters ($\alpha, \beta, \delta, \gamma$) to see how that changes your solution.
If you are interested in exploring error bounds beyond the classic model, I'd suggest exploring a stochastic version of the model or a different modeling regime.


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Does my dracaena marginata have spider mites?
I'm trying to determine if my dracaena marginata has spider mites. A little history about the plant: I got it about 5 months ago and it seemed to be an outdoor plant with the previous owner. Since bringing it home with me it now lives only indoors. I noticed some small cob webs as seen in the photos but I think those have been there since I got the plant. I recall seeing those little webs when I got it. As seen in the photos there are some small holes on the leaves but those have been there since I got the plants. I don't think the holes got bigger. I'm thinking the plant had spider mites at one point but they might be dormant now? I don't see any moving things on the plants at all. I sat and looked at a lot of the leaves but haven't seen 1 moving thing on it. This makes me think the plant once had spider mites but now it doesn't. I'm very new to houseplants so any info is appreciated

Short answer: probably not.
Longer answer: Spider mite infestations present with very fine webbing coating whole areas of the plant, generally whole leaves and the stem. Within this webbing you should be able to see small ~1 mm (0.04 in) dots moving, but it does take some observation and helps if you know what you are looking for. You will be able to feel this webbing if it is present, but may not be able to see it, as it is very fine and best seen by holding up against a dark background with a light source shining on the leaves at an oblique angle.
Holes in the leaves are not indicative of spider mite activity - at least not holes of that size. Spider mites are so small that they feed off the contents of single plant cells, which range from 10 to 100 micrometres in size (that's a 100th to a 10th of a millimetre). This kills the cells, leaving a small pale yellowish dot on the leaf, but not an actual hole that you can look through.
A good example of a heavy infestation can be found on the Wikipedia page for spider mites. I've reproduced it here (file attribution: Paramecium, CC BY-SA 3.0 https://creativecommons.org/licenses/by-sa/3.0, via Wikimedia Commons):

In this picture, you can see the webbing between the stem and the leaves, but note the difficulty seeing it on the leaves themselves, even though it is there. You can also (if you look closely) see the damage caused by the mites themselves as the pale yellowish speckles on the leaf to the top right and middle left.


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Ecosystem naming discrepancies between LANDFIRE ESP and GAP/LANDFIRE National Terrestrial Ecosystems 2011 Datasets
first post here! I am working on a project using the LANDFIRE ESP (environmental site potential) categories, and saw that on NatureServe Explorer I could find broader classifications for the categories provided in the ESP data like this example for the rocky mountain subalpine mesic-wet spruce fir forest and woodland:

I found a table with all of these different levels of classification from the GAP/LANDFIRE National Terrestrial Ecosystems 2011 Dataset, but some of the ecosystem names from LANDFIRE ESP data do not match with any from the GAP dataset. for example, if you search for ecosystems with 'riparian' in the name you get pretty different results from both sets:
(GAP Dataset):
array(['northwestern great plains riparian',
       'western great plains riparian woodland and shrubland',
       'central appalachian riparian   forest modifier',
       'central interior and appalachian riparian systems',
       'ozark ouachita riparian',
       'south central interior small stream and riparian',
       'southeastern great plains riparian forest',
       'mississippi river floodplain and riparian forest',
       'mississippi river riparian forest',
       'southern piedmont small floodplain and riparian forest',
       'edwards plateau riparian',
       'columbia basin foothill riparian woodland and shrubland',
       'great basin foothill and lower montane riparian woodland and shrubland',
       'northern rocky mountain lower montane riparian woodland and shrubland',
       'rocky mountain lower montane riparian woodland and shrubland',
       'rocky mountain montane riparian systems',
       'rocky mountain subalpine montane riparian woodland',
       'north pacific lowland riparian forest and shrubland',
       'north pacific montane riparian woodland and shrubland',
       'california central valley riparian woodland and shrubland',
       'mediterranean california foothill and lower montane riparian woodland',
       'mediterranean california serpentine foothill and lower montane riparian woodland and seep',
       'north american warm desert lower montane riparian woodland and shrubland',
       'north american warm desert riparian systems',
       'north american warm desert riparian woodland and shrubland',
       'tamaulipan riparian systems',
       'rocky mountain subalpine montane riparian shrubland',
       'north american warm desert riparian mesquite bosque',
       'introduced riparian and wetland vegetation'], dtype=object)

(LANDFIRE dataset):
array(['inter mountain basins montane riparian systems',
       'north pacific lowland riparian forest and shrubland',
       'north pacific montane riparian woodland and shrubland',
       'rocky mountain montane riparian systems',
       'rocky mountain subalpine/upper montane riparian systems',
       'california montane riparian systems',
       'california central valley riparian woodland and shrubland',
       'north american warm desert riparian systems',
       'central interior and appalachian riparian systems',
       'gulf and atlantic coastal plain small stream riparian systems',
       'tamaulipan riparian systems', 'edwards plateau riparian'],
      dtype=object)

Any clues on this discrepancy or where I could find a lookup table that better matches the LANDFIRE ESP ecosystem categories? Thank you!!


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Is this a wandering spider?
I recently visited the Peruvian Amazon near Puerto Maldono and found this spider. Can someone please help me in identifying it? It was about 7cm in length, including legs.
I hope that it might be a wandering spider, but idk maybe it could be a massive wolf spider or some other spider…?
I poked its leg gently with a long piece of grass and it didn’t flee, but raised it’s front two legs (only slighly though) as I’ve seen wandering spiders do in videos, and I’ve never seen wolf spiders do this?
Idk if this helps, but anyways, would love some help!

Nice! It's not a wolf spider; the eyes are wrong for that. I think it is; the single dark line on the dorsal carapace and the striped palps certainly suggest Phoneutria, but I'm not sure, so this is a comment, not an answer. It may not have felt threatened enough to go full-on upright. I hope someone else knows.

Yes.

Ctenidae, number 7 on this graphic, have a distinct eye pattern from the wolf spiders (lycosidae; number 1).
I couldn't tell you the species, though.

Awesome, thanks so much for the help!

Just in case you're curious, the largest wolf spider that you'd likely find in Peru is probably Lycosa erythrognatha, which doesn't get much more than 4-5ish centimeters across.


text:
Why did a resazurin broth bottle turn pink immediately after being inoculated?
I have lawn cultures of aerobic bacteria kept at (2-8 °C), which I am trying to revive since they were kept longer than recommended. First, I tried to incubate a broth culture inoculated with a loopful from such cultures and then subculture the next day onto an agar medium. However, they showed no growth on the solid medium.
I resorted to another approach: I immersed a very heavy growth in a few mls of broth which was inoculated into manual blood culture bottles (rubber capped vials), as those:

Some of the bottles turned pink immediately after inoculation. Subsequently, I stained the immersion of these with simple stain (Safrinin), and it revealed the presence of the targeted organisms.
Is there any explanation for this other than that there were viable cells in the inoculum added to the blood culture bottles?
If there were viable cells, how come the color appeared too soon? Large number of viable cells maybe?
Did I push oxygen in with the inoculum through the syringe, so that the indicator got oxidized?

Some other manufacturers say this for the quality control part "color: Light straw coloured solution with upper 10% or less medium pink on standing", i saw some of the unused bottles have a slight pink color and I thought they were ill-manufactured. Maybe i should do a positive control and a real sterile control (inject sterile broth into a bottle, not just use a non used bottle), sorry I'm a dumb brick.

Definitely follow up with those controls, but very likely introduced oxygen when inoculating the tubes. Presumably there's oxygen in the broth and the syringe. I work with a lot of strict anaerobes, and I'm amazed at how sensitive resazurin indicator is. I can be pretty confident that my culture tubes were left open when cycling them into the anaerobic chamber, but sure enough there's always one or two where the anaerobe media goes pink as soon as I dispense it, just from that tiny bit of oxygenated atmosphere left in the tube.

@MikeyC they deleted my reply for unknown reasons. I wanted to say thank you and if this is not good to fit as an answer then what is!

Humble Mikey, i noticed that the "pink" i was refering to is the same color that appears when i injected mere air into a bottle, although i wasn't using aseptic manners, but if the indicator is too sensitive even to few oxygen, the few cells of air contaminants i might have introduced won't produce that few oxygen, right? Thanks.

i meant to say "..... when i injected intentionally mere air....."


text:
There are plants and animal genera with the same Latin name. What are the scopes of uniqueness?
The genus Leptochilus occurs as a genus of wasps, and of a genus of ferns.
What are the parts of the phylogenetic tree where uniqueness is required, and how many of them are they?
(Where can I read about this, without reading all the individual standards?)
Ref: https://www.inaturalist.org/observations/154939850

Does this answer your question? What instances are there in which two species share the same binomial name?

Thank you! Yet no. If the person who asks is correct, an alternative formulation of my question is: What kingdoms exist with respect to nomenclature?
Obviously plants and animals, but what about fungi - different from animals as witnessed by Asterina gibbosa, but are they among the plants? What about diaphoretickes? Bacteria? Viruses? I only find special articles, no overview.
Ah, and I assume that uniqueness is already at genus level, not binomial. (Both is interesting of course.)

@Falko https://en.wikipedia.org/wiki/Kingdom_(biology)

Ups. Diaphoretickes contains plants of course. I intended to mention Stramenopiles.

@BryanKrause Yes, so we have n kingdoms, where n = 2 ... 8 :-)

@Falko there are few instances where a systematist might confuse an animal with a plant etc., so uniqueness should only need to extend to kingdom.

Here https://biology.stackexchange.com/a/73452/63850 Karl Kjer mentions, that animals, plants, and bacteria have their own independent nomenclatural codes. This seems to settle it. (I'm still reading though.)


text:
Autotrophy vs. Heterotrophy in Chemotrophs
What's the difference between chemoautotrophs and chemoheterotrophs? They do both get their energy from high-energy inorganic particles, right?

"Auto" vs "hetero" in general refers to if they can generate their energy independently or require the consumption of another living thing to generate their energy Introductory Biology, 2.18. In the case of chemotrophs, they gather energy from chemical reactions that are not photosynthesis. The auto & hetero here refer to their ability to synthesize organic molecules on their own, instead of energy Microbiology, LibreTexts 5.1B.


text:
RMSD / Tm metric for comparing local molecular structure similarity?
I would like to compare two molecular (protein) structures, but rather than comparing the total, global structures against themselves to yield an RMSD score (which would be straightforward with a structural alignment in PyMOL or similar), I instead am interested in a particular localised structural region in each protein.  Specifically, all atoms within a 20Å radius of a particular residue's alpha-carbon coordinates.
For example, I would have the local structure of L100 (in protein A) and L200 (from protein B), defined as the atoms that are within the radius threshold of those residues respectively.  To compare the "local structures" of L100 and L200, I know that they should be superimposed at the same site as they are the same residue (so I can "enforce" that the alpha-carbon coords of both residues should be in the same spot, i.e. the origin (0,0,0)).  Now, this becomes a "rotation optimisation" problem in order to get the minimal RMSD score.  I have so far found the Kabsch algorithm as one way to achieve this, as there is no need for working out the "translation" of the two separate structures (again, we know that we should enforce the centre residues to be in the same position).
However, the Kabsch algorithm can only work on the same number of points in structure A and structure B; but I may be comparing pairs of structures with an arbitrary number of atoms.  Additionally, due to differing amino acid compositions in the local structure, a different number of each element may be present between the two structures.
I am not sure if there is a solved way of getting the "distance" score between two local structures in this way; and I also am struggling to understand how other structure scores such as a global RMSD (between two different proteins of different length) or Tm (from Tm-align) work with two sets of points with different sizes.  Would appreciate any help!
TLDR;
How to find RMSD score / optimal rotation between two sets of atom coordinates (like Kabsch algorithm), when the two sets have differing numbers of points?

Don't have time to look into it too much right now but have you looked at methods like this https://bmcbioinformatics.biomedcentral.com/articles/10.1186/1471-2105-7-40


text:
Does sunlight interfere with the efficiency of compound light microscope?
I have access to an olympus bx41 microscope. The table on which sits the microscope is in front of a large window and unfortunately no curtains are available. I'm starting to relate that whenever the sunlight is strong, the microscope picture appears dim.
Does sunlight interfere with compound light microscope?

I'm not sure that this is exactly biology, more physics, however the answer is sometimes, though this depends on the type of microscope and how the light is impinging on it.
This answer assumes a common upright compound microscope.
The reason I say this is that microscopes used to (and some still do, such as this one. I have no affiliation with the linked site nor is this an endorsement of the site) have mirrors for reflecting the sun so that you could see your sample.
However, what you are seeing is that your samples appear dim when there is bright light in the room. What is actually happening here is that your eyes are compensating for that level of brightness in your peripheral vision, constricting your pupil so that less light enters the eye. This is called the pupillary light reflex and happens every time your eye is subjected to light. Because of the constriction of your pupil, the central (microscope) image appears dimmer.
Having said that, bright light entering the condenser, not from the (diffused) point-source of the bulb can cause loss of image definition because the light source is not linearized and condensed fully by the condenser arrangement. This results in some images being a bit less sharp. Nikon's MicroscopyU has some detail on how this works.

I actually forgot to say that my eyes + phone pictures both appear dim. If it's the last case then its the condenser that i need to protect for intruding light.

Any idea of how could this be possible?

@FreezingSoul It would depend on the exactly how the phone is taking the images, but light leakage might pay a role in that. You would need to set the shutter speed and aperture for it to take a consistent photo of set brightness, not rely on automatic mode. It's as simple as creating a shade for the microscope - a piece of card taped to the window would work...

i forgot to mark this is as the accepted answer for which it is an accepted answer. i hope i get an illness for this mistake.

@FreezingSoul no need to feel bad - as you might have noticed, most questions with answers don't receive an accepted answer mark.


text:
What determines whether a cell divides by Mitosis or Meiosis?
What I learn from biology class is that mitosis is for producing somatic cells in animals and the gametes of some plants. And meiosis is for production of animal's gametes.
I am curious about what determines whether it divides by mitosis or meiosis.
What signal does it need before it starts mitosis or meiosis?


text:
What kind of tree has this blossom?
I filmed this beautiful blossom on a tree, this past spring in Toronto, Ontario, Canada.

Is it a small tree? It's clearly in the rose family, but that includes apples, pears, quinces, apricots, plums, cherries, peaches, raspberries, blackberries, loquats, strawberries, roses (duh), hawthorns, almonds and more. Best guess? Some kind of plum, like (but not) Thundercloud Plum, because the leaves are out as well as the buds, and the leaves will be burgundy colored. It might also be a crabapple, likely Crabapple "Royalty", but the bark is off. It would help to have a better picture of the leaves and bark, Not an answer because I'm guessing.

Woah, just saw your other posts. You did this last year; you posted a bunch of photos you took in a botanical garden then asked for their names. It's one thing to be genuinely curious about a plant you've come across; it's another thing altogether to take a bunch of photos and ask someone else to identify them for you so that you can label them on your site (or whatever.) -1.

Hello, thanks for your guess and the intel anongoodnurse. It is a fairly large tree in my neighbourhood. Probably 25 feet high, 20 circumference.
Re. your other comment, I shoot stock footage so i'm really particular about describing what i have shot accurately. I am also genuinely curious about this beautiful world of ours that we live in! I also don't think it hurts to increase knowledge on stack exchange via these pics? For example, I posted a pic of a beetle a while back, which prompted a really great and interesting discussion that many people benefitted from, including me!

Maybe, maybe not. I think you're asking others to do work that you should have done yourself at the time you took the photos.


text:
What software can be used for simulating human meiosis, taking linkage and recombination into account?
I want to study the nature of genetic variation in offspring from the same set of human parents. To this end, I would like to take two (male and female) complete genomes, generate gametes from them, combine pairs of them at random, and analyze the distribution of the resultant offspring’s genomes (calculate variance of polygenic risk scores, etc.).
In the absence of recombination, this is trivial; I just duplicate gamete DNA and segregate the vectors representing tetrads. However, I want to take recombination and linkage into account. Of course, I could randomly generate crossover sites and even encode interference in the algorithm I write to perform this, but I want things to be as similar to observed human recombination dynamics as possible.
I can’t seem to find any software packages (preferably in Python, but I’m not picky) that would allow me to do this, but I figure I must have overlooked a standard library for this purpose as this seems like something people would have wanted to do long before now.

I believe that this is an application of the ms program, which is widely used in population genetics.
More recent programs also exist (here is another; here is another, here is another) that may be more applicable directly to human genomes.
I'd suggest looking at those various options and deciding which of them fits your application. This is certainly not exhaustive, there are plenty more!


text:
Did operon evolve after endosymbiosis and specialisation of mitochondria?
We know that all extant bacteria use the operon system as their mode of regulating gene expression, in contrast to eukaryotes, which use individual gene promoters as well as epigenetic mechanisms, supporting the idea that the first eukaryotes came from archaea as the 'host' in endosymbiosis, not bacteria [2]. However, the endosymbiont bacteria is suspected to be a bacterial phylum, supported by various lines of evidence: in short, an archaea enveloped a bacteria to make a eukarya.
An obscure group of protists was found to have mitochondrial DNA that used operon [1], but other than that, pretty much all eukaryotic cells have mitochondria whose mtDNA does not use operon, instead using similar regulatory mechanisms to eukarya including some basic epigenetics (methylation) [3]. A few operons, mainly for nuclear rRNA genes, have been found in chordates. But clearly it is not the norm.
Based on this, it would seem that operon evolved in prokaryotes some time after the endosymbiosis event, since the two are present mostly exclusively in independent domains of life. But recent research finds that operon seems to be fairly primitive (if I am interpreting [4] and [5] correctly) and could have arisen easily as a de novo insertion sequence, suggesting it would have come about earlier.
Does anyone have any expertise on whether this is the case? Did operon or mitochondria come first?
References
[1] Gray MW. Mitochondrial evolution. Cold Spring Harb Perspect Biol. 2012 Sep 1;4(9):a011403. doi: 10.1101/cshperspect.a011403. PMID: 22952398; PMCID: PMC3428767. https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3428767/
[2] Gautam Dey, Mukund Thattai, Buzz Baum, On the Archaeal Origins of Eukaryotes and the Challenges of Inferring Phenotype from Genotype, Trends in Cell Biology, Volume 26, Issue 7, 2016,
Pages 476-485, https://www.sciencedirect.com/science/article/pii/S0962892416300022
[3] Sharma N, Pasala MS, Prakash A. Mitochondrial DNA: Epigenetics and environment. Environ Mol Mutagen. 2019 Oct;60(8):668-682. doi: 10.1002/em.22319. Epub 2019 Aug 6. PMID: 31335990; PMCID: PMC6941438. https://www.ncbi.nlm.nih.gov/pmc/articles/PMC6941438/
[4] Yuki Kanai, Saburo Tsuru, Chikara Furusawa, Experimental demonstration of operon formation catalyzed by insertion sequence, Nucleic Acids Research, Volume 50, Issue 3, 22 February 2022, Pages 1673–1686, https://academic.oup.com/nar/article/50/3/1673/6511971
[5] Marco Fondi, Giovanni Emiliani, Renato Fani, Origin and evolution of operons and metabolic pathways, Research in Microbiology, Volume 160, Issue 7, 2009, Pages 502-512, ISSN 0923-2508, https://www.sciencedirect.com/science/article/pii/S0923250809000539


text:
Mesh formation in the wells of SDS PAGE. How to solve it?
I casted 10% resolving gel and 4% stacking gel and every time a mesh like structure ( polymerisation) in the wells are formed.How to avoid this from happening and how do i get clear wells? If someone has any idea about it, it will be a great help. Thank you

Have you tried rinsing the wells with buffer? Do you have an image of this problem? How old are the chemicals you use - especially TEMED and APS?

Is the comb the right thickness for the spacer you’re using?

This issue is driving me crazy too. Although I haven't identified the exact cause, here are the steps I've taken to minimize mesh formation:
(1) Prepare your gel on the same day you plan to perform SDS-PAGE. Do not store your gel in the fridge overnight.
(2) Allow the stacking gel to harden for a longer period (>40min). I suspect that the mesh forms when the stacking gel hasn't hardened sufficiently.
(3) Switch out the comb. It's possible that mesh formation is associated with a specific comb. Mark the comb number and monitor any mesh formation.
(4) Prepare more gel than you actually need. In my experience, some gels with mesh formation occurred while others did not. By making a few extra gels than needed, I was able to select the gels without mesh formation for my experiments.
All these steps are based on speculation; nothing has been proven yet. Good luck and please share your experiences if you get any improvements!


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Does it make sense to calculate the phospho protein (serine) : phospho protein (threonine) of the same protein in western blot?
I want to indirectly measure the nuclear accumulation of DARPP protein through phosphorylation. Their nuclear transport is controlled by phosphorylation of Threonine 34 and Serine 97 residues.
S97 is then dephosphorylated while T34 is phosphorylated as it moves into the nucleus.
So I want to determine cell localization indirectly through the ratio of Serine 97 and Threonine 34.
I know that the ratio of phosphoprotein to total protein is often estimated by immunoblot analysis. I have antibodies specific to each of these phospho-proteoforms of DARPP.
Does it make sense to calculate the phospho protein (serine residue) : phospho protein (threonine residue) of the same protein? Has anyone ever done this kind of analysis? I cannot find the reference anywhere. Thanks!

What exactly are you trying to find out? It makes no sense at all if you don't tell us.

Are the serines and threonines variants at the same position? are they at different positions? How are you imagining distinguishing between them? Do you have p-T and p-S specific antibodies?

@David I wanted to indirectly measure the nuclear accumulation of DARPP protein through phosphorylation. Their nuclear transport is controlled by phosphorylation of Threonine 34 and Serine 97 residues. When S97 is dephosphorylated and T34 is only phosphorylated, it moves into the nucleus. So I wanted to find out indirectly through the ratio of Serine 97 and Threonine 34.

@Maximilian Press They're at different position. One is T34, and the other is S97. And yes, I have specific phospho- antibodies for both.

editing your question to add this crucial information. Some of it doesn't make much sense to me so please edit my edits as needed.

@MaximilianPress Thanks for adding the information! Nobody knows the answer tho..

@oklu most people probably looked at it when first posted and evaluated it as too vague, and never clicked on it again. Someone likely knows the answer but you may need to wait until more people look at it. In the meantime you could try to do google searches on your own.