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Why don't animal cells have both mitochondria and chloroplasts?
Ancestral eukaryotes had both mitochondria and chloroplasts, why do animal cells only have mitochondria. Why did evolution allow animal cells to give up such a great circle of energy? Where the waste of mitochondria contributes to photosynthesis and vice versa. Is this to the weakness of the animal cell membrane, which cannot hold as much water as animal cells? Though oval-shaped blood cells in camels allow them to consume large amounts of water since the cells are more elastic and can change shape more easily. So why? Just curious. Or is there something I’m missing.

Welcome  Christian. This is not simply a Q&A site, here we are expected to show the results of our own investigations into a subject here, you can [edit] to tell us what your searches found and where the sticking points are. You should also take our [tour] and refer to the [help] for guidance as to the ways of any site you post on, see our section on how to ask.

"Ancestral eukaryotes had both mitochondria and chloroplasts". Please clarify and justify this statement with a reference. Are you talking about the same ancestor, and ancestor to what? I know of no evidence that animal cells ever had chloroplasts to "give up".

I’m voting to close this question because it is either under researched or extremely unclear — it appears to be based on an incorrect assumption about eukaryotic evolution. Please do some reading on this subject and then either [edit] or delete this question based on the other users suggestions.

As evolution progressed and still is on the run, heterotrophs, i.e., the non-photosynthetic organisms did no longer need the endosymbiont cell organelles except mitochondria for their energy requirements. As a result, chloroplasts evolved to be vestigial and eventually non-existent. But exceptions do occur in nature. Chloroplasts can be actually found in non-plant eukaryotes. This phenomenon is called kleptoplasty, where an organism can generate energy from the chloroplast even without performing photosynthesis. For further read, check this: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4517925/

From How do I write a good answer? in the [help], Answer well-asked questions. Not all questions can or should be answered here. ——— Please consider deleting this post as it is encouraging the posting of questions that are not appropriate for this site — please see the [tour] and other [help] pages such as [ask] and [answer] for more information. ——— In addition, your answer is almost certainly wrong — I encourage you to read your linked paper more carefully and consult other sources to correct this misapprehension.


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Amino Acid mutation profile for human coronavirus: Why is the mutation from T to I so frequent?
Still watching the emerging lineages of SARS-CoV2 I noticed that the amino acid mutation from threonine to isoleucine seems to be particularly frequent. Counting mutations in a lineage with a lot of mutation (I choose BA.4) kind of confirmed my anecdotal impression: Indeed, T-to-I mutations were the most frequent ones.
They are more frequent than alanine to valine mutations, and the difference in frequency is not explained by a difference in the frequency of the amino acids in the virus proteines (Threonine is somewhat more frequent than alanine, but not that much).
In part this phenomenon can be explained that a T-to-I mutation involves a nucleotide change from C to T which is the most frequent nucleotide mutation of Coronaviruses in humans. However, in order to be fixated in a lineage, the T-to-I mutation needs also to provide some evolutionary advantage. It seems to be a generic advantage of that kind of mutation.
So my question is: What are evolutionary advantages of T-to-I mutations of SARS-CoV2 in humans?

To elucidate an evolutionary advantage, you'd need to consider the structural context each mutation site along with other nearby mutations. The T->I mutation changes the position from polar to non-polar, which can change how that region of the protein folds (if it folds). Thr is also a site for post-translational modifications like phosphorylation or O-linked glycosylation, either of which could be relevant to just about any aspect of infection like receptor binding, cellular entry, regulation of cellular processes, structural stability of the capsid, or immune system evasion.

The change from C to T (or U in the case of RNA) can happen via the oxidation and deamination of the Cytosine (see reference 1 for the explanation of the mechanism).
The mechanisms looks like the following figure (from the reference 1):

The changes the genetic of the codon from AC* to AU* the first coding for Threonine (T) the later for Isoleucine (I) (image from the Wikipedia):

This does not mean that there is any driving mechanism for this mutation or any particular advantage. As the article states there are quite a few oxidative events per day, since a viral infection causes the generation of a lot of viral particles, this reaction will simply happen by chance.
Besides this mutation happening by chance there seems to be a mechanism called the "APOBEC-mediated RNA editing" which seems to edit the RNA of viruses. According the reference 2 this can be observed on other RNA viruses as well, leading to a higher prevalence of this mutation as it should occur naturally. The author speculate about this as a potent driver for diversification and evolution.
In DNA this error is not so problematic since there are two strands and error correction mechanisms but for a single stranded RNA genome every mutation will persist.
Reference:

Oxidized, deaminated cytosines are a source of C → T transitions in
vivo
Extensive C->U transition biases in the genomes of a wide range of
mammalian RNA viruses; potential associations with transcriptional
mutations, damage- or host-mediated editing of viral RNA

This is a great answer (+1) exhibing the mechanism behind the mutations, but it does not explain the prevalence of T-to-I mutations over A-to-V mutations.

@jk-ReinstateMonica I think the prevalence of this mutation comes from the APOBEC-mediated mechanism. Why this is in place is unknown I think. I will try to see if I find some further information on it.

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Does any brain-computer interface model draw on neural temporal codes?
I am studying brain-computer interfaces (BCIs) and neural encoding/decoding for a class. Most decoding algorithms that I have encountered for BCIs tackling movement problems seem to assume a neural rate code (having done my research on this, I include this link, a chapter by Yu and others, where the model on pp. 226, among others, assume that all the information necessary to decode neural activity is the number of spikes in a given time period, as would be the case if the neurons were using a rate code).
My question: does any BCI decoding algorithm (for movement) assume that neurons follow a temporal code?

Have you found BCI decoding algorithms based on machine learning? As those algorithms should be "model free" would be a nice approach to check the relative importance of rate v/s spike coding in the brain.

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How can this coral survive with little sunlight?
Forgive me if this question is off topic, but I’ve seen similar questions so I think it should be on topic.
I’m at the beach currently, and I was looking in tide pools for interesting things. I found some coral in a one.

I don’t know exactly, but I think there are around 6 or so polyps in the tide pool.
It is very possible that I have misidentified these organisms as coral, when they are something else. If I have, let me know.
It is my understanding that coral survives with the help of algae, which produce food for them. However this tide pool barely gets sun, and is the only one I have found with coral.
Here is a (bad) drawing of the location.

As you can see, the amount of time that sun reaches this location is very short, so wouldn’t the coral be unable to thrive with little sun?
Image for scale of a dead coral (each polyp around 3mm) next to the found “coral”

Welcome to Biology.SE. I can't tell what it is in your picture that you think is a coral. Can you supply a clearer pictures? If not, please indicate (e.g. with an arrow or circle) the relevant part of the image and add a rough sketch of the organism(s). In addition, for species-identification we need to know the size, and geographic location. That said, coral polyps are typically very small — is it possible you are looking at sea anemones?

@tyersome It’s basically dead center of the image, as well as another one on the far bottom right. I guess calling it coral is an overstatement, it’s very few polyps. I don’t have a clearer picture, but tomorrow I could try to get one. it could be hard to tell scale, the size is about 4mm. I don’t know exactly though, it’s night and so I’m not going to go looking for the spot right now. I am currently on the coast of South Carolina. It is definitely possible that it is a sea anemone, however it is vastly different from the other sea anemones i have found in the area, it’s much smaller.

Certain species of corals do not require as much sunlight as others to survive. The zooxanthellae that augment nutrient supply to the polyps do rely on photosynthesis, but some species are adapted to less direct sun sources. For example, there are corals within the mesophotic zone of the oceans where solar radiation is low compared to that of a tidepool (even if it rarely receives direct sunlight). For your specific question, the corals likely receive enough sunlight to meet their energetic demand of symbiotic zooxanthellae to complement their predation feeding strategy.


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What are the apomorphies of Streptophyta that delineate them from Chlorophyta in Viridiplantae?
I'm reading through Plant Systematics, 3rd Edition by Michael G. Simpson, and am having a hard time understanding the differences between the Chlorophytes and Streptophytes - discussed in Chapter 3: Evolution and Diversity of Green and Land Plants.

The cladogram (from the second edition) makes it seem like the development of oogamy and plasmodesmata delineate the Streptophytes from the Chlorophytes, yet, still we see a few lineages within Streptophytes without either of these apomorphies (even in the third edition's version of this cladogram).
Where do we really draw the line between these sister groups?

Welcome to Biology.SE. Don't know the answer, but a quick look at the relevant wikipedia pages suggests they may be helpful — the short version is it depends on how the groups have been defined ... Also note that from your diagram it seems clear that neither oogamy or plasmodesmata are relevant — they distinguish Charophytes from other Streptophytes.

@tyersome the figure in 3.1 of the third edition actually doesn't even indicate Charophytes, but does include a Charophyceae lineage diverging from the rest of the Stroptophytes directly after the development of oogamy and plasmodesmata. Confusingly, Simpson goes on to discuss apomorphies of the Charophytes and references that figure even though there are no Charophytes. This may be an error? I've gotten a hold of the article that this cladogram was roughly based on (https://bmcecolevol.biomedcentral.com/articles/10.1186/1471-2148-14-23), so I'll update this with whatever I find.

@tyersome looks like the evidence for the division between these two groups is primarily genetic data. I will answer my question below shortly.

In the article cited by Simpson in creating the rough cladogram of Figure 3.1, From algae to angiosperms-inferring the phylogeny of green plants (Viridiplantae) from 360 plastid genomes (Ruhfel et. al, 2014), the phylogeny of Viridplantae is inferred from a set of 78 genes.
Thus, the division of Viridiplantae into the two sister groups Chlorophyta and Streptophyta is inferred / supported by molecular evidence.


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Photo attached of flies ‘injecting’ into spider egg sacs. Why? What’s going on here?
Brisbane, Australia.
While no image is available, direct observations were made of flies doing something to the egg sac.

Welcome to Biology.SE. What makes you think the flies are 'injecting' something into the structure? Also please give us any information you have relevant to species-identification — in particular it could be helpful to know the geographic location as well as the ids of the species involved (if known).

Possibly cannot see in the photo, but I could see each fly had a part of themselves extended from their rear end going into the egg sacs.        She is a ‘tent spider’ and that’s how they lay their eggs. Each ‘bump’ is a sac of eggs.

Cyrtophora moluccensis- tent spider.
Injecting? I could see something from the rear end of the flies going into the sac. (Sorry, Not clear in photo).                                        Location: Brisbane Australia

(Converting from comment)
Based on internet searches, it appears likely that the fly is Sarcophaga arachnivora, which lives off of spider egg cases in its juvenile form; the fly is laying eggs.
If you follow that link you will see references to a couple of papers that describe the species in more detail, though I don't believe I have access to them.
The geographic location noted there of SE Australia agrees with your observation of the organism.
References about tent spiders also make note of parasitic "flesh flies" so the species interaction appears to be an established phenomenon.


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Resource to obtain listings of tissues that different viruses infect
I am working on a project that involves analysis of viral DNA sequences. I'd like test hypotheses about issue specificity. Ideally I'd be able to get a table of virus/strain names with a column in it for something like 'tissue tropism' which would equal neurons/nervous system, liver, epithelia, etc etc. Does anybody where I could find this?
I so far haven't located it www.viprbrc.org which seems to be molecular bio info like domains, dna sequence etc.
I also found this paper paper which has data in the ideal format but is limited to analysis of RNAseq data and to healthy subjects.

I understand the question, for example HPV targets skin cells, HBV liver, however a lot have complex/cryptic lifecycles for example the different HHV. My view of this information/question is that it is a classic virology textbook rather than a singular database. DNA viruses are quite a specific group, which includes VZV (chicken pox virus HHV3) and smallpox BTW. They often form complex chronic infections (HPV many HHV) and thats not really database stuff. Transferring  this question to biology.stackexchange is probably the right place for it IMO.

Tried viralzone?

Animal viruses? Vertebrate viruses? Mammalian viruses? Human viruses? Please clarify.


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Vegetation density estimation from distance method: random sampling or along the grid?
I am planning to sample the tree regeneration (new ones and advance regeneration, > 2m). I am planning to use small circular points to estimate the counts per area (= density of young trees). I wonder about the sampling design: what is better for following whole area tree density estimation? If the sites would be placed randomly, some might overlap. Is the sampling across the grid a better option, but requires a set number of subsamples: eg. not very scalable?
For any of the metrics, we will then calculate distances to the closes tree. Is any of the metric better in folliwing density estimation from teh distance methods?
Or, what I found, a quadrat sampling, combining both aproaches. Seems however quite challenging to do in the field. What are your thoughts? Thank you.


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How to look for different transcripts for a protein?
I am trying to find all the different transcripts for a protein that translate into different isoforms of the protein.
However, when I look it up online, there is no clear organized data on different documented isoforms (at least not for the one I am looking for...) (For example, entries at NCBI Protein database do not match with those at ENSEMBL).
It would be great if someone can suggest a more systematic way (or previous such studies) to find out the sequences of different isoforms of a protein.
To elaborate, here the link at ENSEMBL database (9 transcripts, of which 5 are protein coding, of which 2 have identical AAs seq) and here's what I found at NCBI Protein search (4 isoform and a 5th one from literature). It is evident that there are discrepancies between the two.

Are you looking for isoforms or just differential transcripts? Isoforms can be produced from completely separate genes that have undergone duplication at some stage. 2. We need to know what gene you are interested in and in what tissue. Don't be coy. If you find differences between ENSEMBL (no E) and NCBI there must be a reason — both are reputable sources, but may be providing different info. I don't see going to the UCSC browser resolves that. Tell us how the info on the two sites differs and give a link.

I think David's first question bears repeating - are you looking for varying transcripts from the same gene, or related proteins with high sequence homology from different genes?

Hi!

I am looking for varying transcripts from the same gene.
I have also added the links to my search from both the databases. Hope you can see the discrepancy I am facing.
Please let me know if I have missed/misread something here.
Thank you!

I've revised my response to your question, but I think you will find that there is no easy "answer" to it. If you are involved in practical research that depends on your analysis of this question, you are going to have to do some serious reading before investing time and money.

As will be clear from the question and various replies, there are several different repositories for data emerging from the various nucleic acid sequencing projects. The two main gene oriented ones — NCBI and ENSEMBL — are funded by the US government and European goverments, respectively, and share primary sequence data. However they differ in their interpretative focus and the secondary data (often from elsewhere) that they present in combination with the nucleic acid sequences. There are other valuable webs services that are more limited in scope — the UCSC browser (mentioned by Maximillian Press) providing graphical views of several reference genomes. The main protein-oriented repository is UniProt, as @Michael_A mentioned.
The poster is interested in the transcripts for a particular gene (Transmembrane protease serine 3), and the NCBI and ENSEMBL facilities he consulted were, in my opinion, the most appropriate. As he found different transcripts presented by the two facilities, he might follow the suggestion of consulting UniProt, but it transpires that the set of isoforms it presents differ again. (The UCSC browser displays the ENSEMBL set which originate from the GENCODE project.) This is summarized in the table below — three common isoforms, but ones absent from other data repositories.

NCBI
ENSEMBL
UniProt
Size (aa)

NM_024022.4, NP_076927.1 (iso 1)
TMPRSS3-203
P57727 (-1)
454 aa

NM_032404.3, NP_115780.1 (iso 2)
—
P57727-2
327 aa

NM_032405.2, NP_115781.1 (iso 3)
TMPRSS3-201
P57727-3
344 aa

NM_001256317.3, NP_001243246 (iso 4)
TMPRSS3-208
P57727-5
453 aa

—
TMPRSS3-202
—
451 aa

—
—
P57727-4
293 aa

AAT66641.1 (iso 5)
—
P57727-6
538 aa

[NM_024022.3 (= NM_024022.4)]
[TMPRSS3-209 (= 208)]
—
[duplicate]

Why could cause this conflict, and what can one do to resolve it? Transcript data may be obtained for individual genes in specific studies, or in large projects like ENCODE. Whereas I imagine the commonalities probably arise from the larger projects, it seems likely that the smaller ones are represented in the individual unique cases. The reason that this might be so is that the pattern of differential splicing often differs between tissues and at different developmental stages, so that such transcripts might be missed in whole body studies or in studies in which a tissue was not included. In this case uneven coverage of tissues involved in hearing could possibly be the problem. Mistakes can also arise: this is presumably the reason for the two identical transcripts being listed. And it is difficult for NCBI to co-ordinate different submissions — hence the strange situation of the missing transcript map for ‘isoform 5’.
So there can be no general answer as to which data repository is the “best” or the one to “believe”. One has to find where the evidence is for a particular transcript, i.e. identify the original papers, read them, and make one’s own decisions as a scientist.

This is almost a complete revision of my previous answer. Apologies, but I think it provides a more balanced view of how things are. However, anyone who voted for the previous one but doesn't like this one should retract their vote.

I would suggest using the UCSC genome browser for the organism in question (assuming that they have your organism loaded), and use either the GUI or otherwise to pull out the transcripts you are interested in after searching the gene. For example, here is a view of transcripts for the CFTR gene in humans.
You can click on the transcripts in the cartoon, leading you to a page such as this that has a lot of information.

As you are interested in protein coding transcripts it would be worth using a protein database. You can search for curated protein isoforms in Uniprot by limiting your search to "UniProtKB" + the species. Searching for "TMPRSS3" shows 6 protein isoforms. The cross-references section of the page provides links to the RNA transcripts for each isoform in a various DBs. A quick scan confirms that there are direct links to transcripts for 4 of the isoforms; if you check a little closer it should also provide a direct link to the other 2.

I think people more or less expect to do BLAST searches, and also to check the literature when it's available, when they really hope to find all potential splice forms that are currently known.  Deciding when a transcript is genuine mRNA rather than partially spliced RNA from the nucleus can be tricky, and may depend on the specifics of how the RNA was isolated (e.g. polysomes vs. whole tissue homogenate; though the difference, while real, seems to be less than expected).  Also, there's a philosophical problem: some of the transcripts from your first link barely overlap with one another, and I don't think it's uncommon to look at a stretch of genome with exons ABCDE where AC, DE, and BCDE are all possible splice products - are they "the same" gene?


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What is the DNA sequences between two Inverted Repeats called?
I would like to know some rules about the nomenclature of Inverted Repeats DNA sequences. More in detail, I want to know what is the name of sequences between two inverted repeats.
For example, I have two genes A and B in this order: A, B, -A. The gene A is inverted duplicated and forms an inverted repeat with gene B in the middle. What do I call this gene B?
Here below are some names I found or that I thought, however, I don't consider them a good choice. Could you suggest some other names?
"intervening sequence", "in-between", "inserted", "separating", "spacer", "non-palindromic spacer", "loop", "gap".
In Wikipedia, they used the name "intervening sequence".
https://en.wikipedia.org/wiki/Inverted_repeat

Welecome to SE Biology. Come si chiama, como se llama, Wie heißt, comment s'appelle. But in English is is What, not How.

There is no "rule" or even convention, as you found from Wikipedia. Such sequences are not sufficiently important (and have no general function) to have been given a name. Intervening sequence is a description, but as it is the long form of intron, I wouldn't use it. For suggestions on how to describe it (there are problems with all of yours), SE English Language & Usage would seem more appropriate. However you will need to give an example sentence there, which should relate to the specific circumstances.

To add to David's points, "inverted repeats" typically described very short sequences. Even so-called "long inverted repeats" are just tens of bp long. Since you have designated sequences A and B to be genes, it would be better to described your system as "gene B flanked by inverted copies of gene A", or something like that.

Thank you David and acvill, both comments were very useful. Yes, unfortunately, I have some long Inverted repeats with many genes and kilobases long which are not common and not well described (named).
I want to name them to describe a property that I think these sequences might have. I think I will use the term "internal sequences" or "sequences flanked by internal repeats" as acvill suggested.

I'll add that most green-plant chloroplast genomes contain a large inverted repeat (~10-76 kb). The chloroplast genome is circular, and the two intervening regions are creatively named the Large Single Copy and the Small Single Copy.


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Alignment given two sequence
I have two sequences and I need to perform sequence alignment to determine all possible sequence alignments. I have created the matrix and managed to find 16 alignments.
I wanted to understand if I had correctly approached this because I need to provide all possible alignment sequences and then provide the best alignment sequence/sequences out of all the ones found, however, all my found sequences have a score of -9. The sequences I have found I have placed in this pastebin link: Sequences since I did not want to make the post too large.
I was wondering if someone could help me understand if I may have gone wrong or if I am missing some sequences since all of them have a score of -9. I do not think I have gone wrong during the matrix creation phase but I am unsure. My goal is to find all sequences and not just the optimal ones.
I have asked for help from my tutor however he has been quite rude and reluctant to explain/help so I have decided to turn to the community for help and guidance. Would appreciate any insight into this. Thanks in advance.
Sequence strings:
ggaatggmeeff
gatge
Alignment matrix

The poster writes “I need to perform sequence alignment to determine all possible sequence alignments”. It is not clear whether this need is that determined by a course assignment or the poster’s mathematical  imagination. If the former, I doubt that the wording expresses the request correctly, even though this is repeated in the question as “My goal is to find all sequences and not just the optimal (highest?) ones”. The reason for doubting this is that the number of possible alignments with no restraint imposed will vary from very large to astronomical, depending on the length of the sequences.
The display presented looks at first sight like a trackback for a global alignment using the Needleman and Wunsch dynamic programming algorithm. Such a trackback is designed to find one of what are generally several possible alignments that have the highest score using a specific scoring matrix and gap penalties (different scoring systems can give different alignments). Although it is possible in general to trace alternative trackbacks of the same score manually, I do not see any in this example. This operation is not performed in any programatic implementations of the algorithm of which I am aware. Although, memory limitations permitting, one could program this, the algorithm is not designed to find lower scores (and I find it hard to see the demand for such).
However the problem with the Needleman and Wunsch trackback is that in this algorithm each cell should trackback to the single highest-scoring cell preceding it — if there are two of the same score a random choice is made. But the trackback shown, as well as having instances where the highest scoring cell has been missed, has 9 cases where the trackback is to two cells with different scores. I can think of no justification for this.
There is another fundamental defect here, which arises from the scoring system used. The match = 1, mismatch = –1, fixed-gap-penalty system is doomed to result in low alignment scores in this case: the best possible score is 5 x 1 (matches) + 7 x –2 (gaps) = –9. The PAM or BLOSUM matrix, which score similar amino acids on the basis of their likely mutational substitution and thus are better related to the realities of protein structure,  have higher numerical values relative to the gap penalty, which should actually be replaced by a gap initiation penalty followed by a lower gap extension penalty.
In conclusion I would advise anyone undertaking work of this type, first to obtain a thorough understanding of the dynamic programming algorithms used, next to specify clearly the goals he wishes to achieve, then to assess whether they are realistic, and finally be prepared to write a program himself if none already exists.

Thanks for the response. We get weekly activities for practice to compliment the lecture. The question states to find all possible alignments, followed by "provide the best alignment sequence/s". However, from my understanding and as you have mentioned, the most optimal alignments are those that go from the most bottom-right to the most top-left. I have managed to find 16, all of which have a score of -9. Since I found them all with the same score, I thought my approach might have been incorrect. My tutor is not polite when you ask for help and so I decided to turn to the community for help.

@Krellex — It's possible for a lecturer to say or write something in a lecture that's ambiguous or not exactly what you intended — I've done it myself many times — and students can sometimes get hold of the wrong end of the stick. It's difficult for someone else to tell. However the generally topic is covered on the web in many places other than the Wikipedia link I gave. Have a look round, and perhaps discuss it with your classmates. Double-check your trackback, and perhaps try a BLOSUM matrix for scoring.


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What bird is singing at almost 3 AM in Hong Kong? (recording included)
I live in Hong Kong and there’s a bird around my street that sings around 3 AM, when there isn’t the slightest bit of twilight.
I have recorded its song here: https://youtube.com/shorts/0dBNvPAgLmU?feature=share
Can somebody identify it?

+1 for providing an actual recording! Can you describe a bit more about the song? For example, is it several seconds long and repeats, or goes on and on for up to a minute in a somewhat rambling, non-repeating way (which describes what I often hear between 3 and 5 AM in Taiwan)


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Is there a specific name for the feet of insects?
A friend and I were talking, and he asked me whether there was a name for insect feet (either in general or for specific insects). I think they are sometimes called tarsi, but I believe that's a more general term.
He is wondering about bees in particular. More specifically, he asked me, "If you were to put handcuffs on a bee, what would you call them?"
If any entomologists here have an answer, we would both appreciate it! Please rest assured that the handcuff question is purely hypothetical; no bees were harmed in the asking of this question!

Welcome A Bee. Please enjoy our [tour] and refer to the [help] as and when for guidance.

Did you know that you can google insect anatomy? On this site, we ask that people try to search for an answer themselves before asking, and to ask for clarification if there's a specific problem.


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Stimulus currents and neuronal responses
As I understand it, if a subthreshold current of unlimited duration is injected in a neuron, a passive response is observed, like an RC circuit. The membrane potential is depolarized by some arbitrary amount, before gradually stabilizing due to more and more current flowing through the resistive branch of the circuit.
So yesterday, I was looking at some HH simulators when I noticed that increasing the duration of a normally subthreshold current pulse would allow it to depolarize the membrane to threshold. How is that? Can someone please point out the error in my logic here?
Also to clarify: I am not familiar with the dynamics of the HH model, so please take it as such.

If you inject a current into a model cell with no active components (no voltage-gated channels), you'll see a capacitive response, because that's basically all you have: the membrane acts like a capacitor in parallel with a resistor. Adding a current charges the capacitor and you have an "exponential decay"-shaped change towards a new equilibrium potential.
When you stop injecting current, you'll again have an "exponential decay"-style return to the baseline.
However, the HH model includes active conductances: voltage-gated channels that change state according to voltage. "Action potential" is the name given to a positive-feedback opening of sodium channels: voltages more positive than rest open some channels, which open additional channels, etc. "Threshold" refers to a voltage that triggers a full positive-feedback activation of these channels. Sometimes it's stated to be a particular voltage, but that's not really true; threshold depends on the whole state of the system: voltage and state of all the gates of different channel types.
However, any depolarization will open some voltage-gated channels beyond those open at rest, it's just that up until threshold there aren't enough open to trigger the positive feedback. The reason is that voltage-gated sodium channels don't just open and close, they also inactivate. In the HH model, this is represented by the "h" gate.
If you inject a brief positive current, you'll open some channels and get a little "bump" in the voltage afterwards that then returns to baseline as those channels close/inactivate. If you inject a longer current, you'll open a few more channels and also the "bump" from the open channels will sum with your injected current. If you're right around threshold, that could cause you to exceed the threshold for positive feedback and therefore an action potential.
Though the question is different, a lot of this same reasoning is found in my answer to another question here: Hodgkin huxley neuron not spiking consistently for currents greater than threshold?

Thank you for your prompt answer! I think I get it now. Can you just allow me one more question? It’s been bothering me a bit!

It goes like this: a stimulus current, at the beginning of an AP, depolarizes the membrane towards threshold right? For some reason textbooks always pretend like they don’t exist once the membrane reaches threshold and an action potential fires. So let’s assume that the stimulus current is on, for the whole duration of the action potential. My question is: would that change any of the AP’s characters? For example, increase its amplitude or duration? The one with amplitude specially resonates with me, as I imagine that the current would depolarize the membrane all the time while it’s flowing.

@CalyxOfHelp Happy to answer if you post as a separate question. Would be ideal if you quoted from your textbook or another citable source, and take some steps to think about what the answer might be and describe your thinking (sort of "show your work"). That'll help me write an answer that specifically addresses the gaps you have.

Ok, I'll try to write it soon, thank you!


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Why does fresh zinc oxide cause chills but "old oxide" does not?
Inhalation of fresh zinc oxide causes chills; aka- zinc chills, brass founders ague, oxide shakes, etc. But if the oxide is a few hours old it will not cause chills. I had them once long ago, an old experience person explained the situation which I had never heard of. Pollution controls and modern safety practice have made it very rare in the US today. It can occur if one inadvertently causes a "zinc boil" by overheating a brass melt or oxy-acetylene cutting galvanized steel, etc.

Can you include a reference to something besides your personal experience? It seems very unlikely to me that anything here is involving the "fresh"ness of anything; it sounds like you're talking about a situation where there are inhalable metal oxides present compared to a situation where there are not inhalable metal oxides present. In that case the "why" is a bit boringly obvious, is it not? Can you clarify your question to make it more clear where the uncertainty lies?

I'd wager that your "old zinc oxide" is nothing more than rough-powdered whereas the irritant stuff from welding is nanometre-sized and finely dispersed in air (from experience and observation).

this is possibly because fresh zinc in the air is not fully oxidized and the old zinc in the air are.maybe you can ask a related question over at https://chemistry.stackexchange.com/ and link to this one.

Is what you're describing the same as metal fume fever?  This article describes it and says the particle size should be under 1 micron for acute systemic effects, and also says that aggregation of these particles occurs.  Do you know whether the old oxide you describe clumps up into larger particles?

Note the reference states "freshly formed" zinc oxide. Also, Zinc oxide is produced by mixing hot zinc vapor ( very small particles) with air. Wikipedia article not accurate ; i am looking for the book reference.


text:
What's the effect of prolonged threshold stimulus currents on action potentials?
I've been confused as to what exactly occurs when a current is injected into a membrane, throughout the duration of an action potential. My main source of confusion has been trying to reconcile 2 things, mainly the invariance of the action potential's magnitude, and the logic that I used during my study of action potentials to explain their origin from equivalent circuits.
To illustrate:

This is a diagram from Cellular Physiology and Neurophysiology, a textbook that I use from time to time. This diagram has been of great usefulness to me, as it provided me with an equivalent circuit description of all phases of the action potential. As you can see during B, a current is injected into the membrane, depositing a net positive charge on the membrane capacitance, causing an outward capacitive current that discharges the membrane capacitance to threshold. Immediately after reaching threshold though, the injected current terminates and the rest of the depolarization is done by the inward Na+ current, which is in a positive feedback cycle.
Now comes my problem: what happens, if I deliberately keep injecting current throughout the whole action potential? Would, say, its magnitude change? An easy answer would be "nothing", as the action potential is always the same regardless of stimulus strength & duration. However, some of my logic is making it seem like the action potential should be changed.
Allow me to explain. My problem comes at the peak of the action potential. This is supposedly a steady state, where the decreasing inward Na+ current is equal and opposite to the increasing outward K+ current. However, for our experiment, you would have to account for the constant inward injected current, which would deposit a net +ve charge on the inside of the membrane. Now comes the inevitable question: wouldn't that increase the peak of the depolarization? For a normal AP, the peak will always be a little less than the equilibrium potential for Na+, however here the presence of the constant inward injected current (and by extension the constant outward capacitive current) makes me feel as if it would nudge the peak a bit higher, before the outward K+ current can eventually eclipse it. Is that right or not? I feel like I'm missing something crucial here.
Another way to put it, in equivalent circuit terms:
. at threshold, Ic = Im + INa
. INa decreases with more depolarization, Ic
gradually decreases
. At the peak of the AP: Ic = Im (my proposed
nudge beyond ENa)
. After that, Ik > Im + INa, and should be able to repolarize the membrane.
So yeah, I guess it all boils down to me not being able to work out what happens in a neuron when a constant threshold current is injected. I'm very sorry if this is too long, and many thanks for reading! I've tried to explain things as best as I can, please tell me if something is not clear.
Diagram source: Blaustein, M. P., Kao, J. P. Y., & Matteson, D. R. (2019). Cellular physiology and neurophysiology: Mosby physiology series (3rd ed.). Elsevier.

Having taught AP mechanisms to undergraduates and graduates for many years, I find the electrical equivalence circuits less useful for understanding the mechanisms. They date back to Hodkin and Huxley and one should recall that they had no idea about what actually happened on the membrane during an action potential. All they could observe was changes in currents and cross-membrane potentials and they had no other way of understanding the processes than through electrical diagrams.
For me the key is realizing that the currents themselves are of little importance. What really drives the AP are changes in membrane permeability of Na and K. The membrane potential at any time is given by the permeability for Na and K (and Cl), as expressed by the Goldman equation.
$\ E_m = \frac{RT}{F} \ln{ \left( \frac{ P_{\text{Na}}[\text{Na}^{+}]_\mathrm{out} + P_{\text{K}}[\text{K}^{+}]_\mathrm{out} + P_{\text{Cl}}[\text{Cl}^{-}]_\mathrm{in} }{ P_{\text{Na}}[\text{Na}^{+}]_\mathrm{in} + P_{\text{K}}[\text{K}^{+}]_{\mathrm{in}} + P_{\text{Cl}}[\text{Cl}^{-}]_\mathrm{out} } \right) }$
where $\ p_x$ indicates the permeability of ion X. If the membrane is depolarized, the Na channels open, which increases Na permeability and draws the membrane potential towards the Na Nernst potential (which is well above positive). When K channels open a little later (together with the closure of Na channels), the potential is pulled back down towards the K Nernst potential, which is below the resting potential. If you continue to inject current during the AP, this is equivalent to introducing a new channel for an ion with a Nernst potential equal to the voltage you apply to the electrode. If you apply a voltage higher than the normal AP potential, the AP amplitude will increase. If the voltage is lower than the normal AP potential, the AP amplitude will be smaller. In essence, you have created a voltage clamp setup, which was the key technology used by Hodkin and Katz when they studied the mechanisms behind the AP, which led to the proposal of the existence of voltage dependent Na and K channels.
A voltage clamp in your electrical equivalence circuit corresponds to inserting an additional battery with a very small serial resistance (equal to a permeability much larger than any of the three ions). This battery now becomes the dominant and determines the membrane voltage.


text:
Any reasonable avenues toward extending telomere length for humans?
Is there any biological research progress toward extending the length of telomeres for humans?
It seems as though this has seen some moderate success in extending the length of life of mice -- is it possible that current living humans could have some way of extending or preserving the length of their telomeres (given some realistic advancement in biological science)?


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How much does each rotation coarse adjustment move the stage in a microscope?
I've been working with microscopes lately and have been trying to think of a way to somehow find out a way to some how find out a way to relate the rotaions of fine adjustment to coarse adjustment.
The main idea is to figure out how many centimeters the fine adjustment moves the stage, then compare the same thing for coarse adjustment.

This is unanswerable unless you specify the particular model of microscope as there are tens of thousands of different models with their own specifications. You would also need to show your research in alignment with the [help]'s requirement. It would be great if you'd also tell us the purpose of the question and how it relates to biology, as it seems to be a question of engineering at present.

The instruction manual for the microscope would help greatly.  But there is a simple method that should tell you what you need to know with modest precision.  Use micrometers (the instrument, not the unit) to measure the thickness of a microscrope slide.  Mark the top and bottom sides of the slide with an expo marker.  Put the slide in the slide holder firmly, and focus on the bottom of the slide first ... then count how many times you need to turn the knob to focus on the top.


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Phenotype-to-genotype mapping
My understanding is that GWAS or QTL attemp to extract potential SNP candidates responsible of a particular trait.
However, is there any framework that attempts to do the opposite? This is, link the phenotypic value of traits to the distribution of allelic frequencies.
For example, given a set of traits construct a classifier (i.e. SVM) for a given SNP and try to predict its allele.

https://en.wikipedia.org/wiki/Phenome-wide_association_study

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


text:
Does HIV erase the immune memory?
It’s said that HIV selectively infects and kills CD4+ T cells, which disables the adaptive immunity because both CD8+ T cells and B cells need CD4+ T cells to activate them. However, once activated, memory B cells and CD8+ T cells can function independently of CD4+ T cell. Does that mean HIV+ people still retain immunity against previously encountered or vaccinated infections even at advanced disease stage?
Cited from:

As the affinity of surface Ig from memory B cells is increased, their
requirements for reactivation are lower than for naïve B cells: memory
B cells may thus be recalled by lower amounts of antigen and without
CD4+ T cell help.

https://www.sciencedirect.com/science/article/pii/B9781416036111500064

Welcome to SE Biology! It would be helpful to link to/cite sources for your information.

Here: As the affinity of surface Ig from memory B cells is increased, their requirements for reactivation are lower than for naïve B cells: memory B cells may thus be recalled by lower amounts of antigen and without CD4+ T cell help. https://www.sciencedirect.com/science/article/pii/B9781416036111500064

Please edit your link into your question - that way it is more likely to be read, and more likely to persist as comments can and do get deleted occasionally.


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Is the transition epithelium a monolayer epithelium? Or is it a stratified epithelium?
I had learned that transitional epithelium is a type of pseudostratified epithelium, so I understood that transitional epithelium is a type of monolayer epithelium.
Wikipedia has the following description(but with source tag);
” These structural elements cause the epithelium to allow distension; however, these also cause the tissue to be relatively fragile and, therefore, difficult to study. All cells touch the basement membrane.[citation needed]”
https://en.wikipedia.org/wiki/Epithelium
From the above description, all　epithelial cell appears to be in directly adhered with the basement membrane.
However, the illustration in the Wikipedia entry for Epithelium is clearly multi-layered. Thus, it appears to be a stratified epithelium.
https://upload.wikimedia.org/wikipedia/commons/thumb/6/64/423_Table_04_02_Summary_of_Epithelial_Tissue_CellsN.jpg/900px-423_Table_04_02_Summary_of_Epithelial_Tissue_CellsN.jpg
So, I would like to identify which is correct;

Is the transition epithelium a monolayer epithelium? Or is it a stratified epithelium?

Urothelium, which is another way to refer to transitional epithelium is indeed a stratified epithelium.


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Is this an Australian Wolf Spider? Any idea on the species if so?
I live in Western Australia, and recently have been finding a lot of a particular type of spider out in my backyard (generally at night, on the lawn).
From what I could find, I believe it is an Australian Wolf Spider, but wanted to make sure by asking here, as I was suggested to do so, when trying to find out how dangerous they are to small dogs.
Here is a pic of one I managed to capture (in a glass pasta sauce jar for size reference), the spiders are about 25mm - 50mm big I'd say.

Here is a close-up:

I have also being seeing the tell-tale sign of small little burrows in the ground for awhile now, which seems to be what they do.
As I understand it, there are many many species of Wolf Spiders, and from this Reddit question a couple of people explain them quite differently to the spider I am seeing, eg;
The wolf spider is big, hairy and scary looking

Wolf spiders, though rather large and fearsome

Is this an Australian Wolf Spider, and if so, any idea on the species?

A closer-up photo showing its eyes clearly would help with identification.

@timeskull I have added a close-up, that's as close as I can get with the current shot. If you need closer, I would need to capture another one and take another photo. Cheers!

This observation might be suitable for iNaturalist.

@AgnesianOperator Interesting. Thanks!

My best guess is Lycosa australicola.
Found across the central and west of Australia.
Other species I have not yet checked that occur there (from wikipedia entry on lycosa, from the World Spider Catalog) are:
ariadnae
corallina
cowlei
dimota
gibsoni
hickmani
koyuga
salifodina
tula
woonda
yalkara

Thanks - though the colouring does seem a bit differently placed on mine?

I haven’t checked the whole list. It might be a new one.

How about you go through the list? If it’s a new species you can name it after me.

Way to many to go through lol

it's an Australian wolf spider, based on:

striped back
long hairy legs
so we can deduce it's an Australian wolf spider.

There are many spiders that fit the description of "long hairy legs, striped back". Your answer could be improved by adding something more specific, with the citation of a reference or two.


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scientific reasons behind a a sudden yet temporary growth sprout of the plant when the soil has No depth?
Sorry, I'm kind of naive when it comes to plant biology.

Mark 4:5-6( New American Standard Bible 1995 )
5 Other seed fell on the rocky ground where it did not have much soil;
and immediately it sprang up because it had No depth of soil. 6 And
after the sun had risen, it was scorched; and because it had no root,
it withered away.

When someone plants in soil with No depth then will the plant immediately spring up( i.e. a sudden yet temporary growth sprout)?  If yes, could someone please explain the scientific reasons behind a a sudden yet temporary growth sprout of the plant when the soil has No depth?

Welcome to Biology.SE. I’m voting to close this question because it is both unclear/unfocused (among other things the type of plant and the environmental conditions will matter) and not well motivated (religious texts are not a good source for biological information). If you wish to ask questions here  please take the [tour] and consult the [help] starting with [ask] for details of what is expected.

Normally a seed would germinate (start to grow) when there is enough damp and warmth beneath the surface of the soil, and the shoot would push upwards at the same time as the root pushes downwards. The root then would be in the moist soil, able to suck-up water when the young plant needs it.
But because there is no depth to the soil, the roots grow and are unable to penetrate the soil and go down, instead they push the body of the seed upwards:

Cress seeds grown on tissue. Copyright unknown, groseeds.co.uk (commercial) 2022 fair usage.
The roots and shoots will then dry out.
You can try this at home with cress grown on moist tissue paper.


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Is it possible to identify a worm being carried by a depicted wasp?
Wikimedia has a picture of a Ancistrocerus trifasciatus / mason wasp carrying a kind of worm:

Is it possible to tell what exactly kind of worm it is, based only on the above image, number of worm's segments, dots on its skin and a correlation in size to wasp's size (or its feeding preferences)?
I am not the author of the picture, I was not present when it was captures, so I can't provide any more details that are usually helpful in species identification (like moment of day and year, geographic location etc.). The image itself doesn't contain any geolocation data, but from its German description in can be assumed that this was captured somewhere in Germany, Europe.

Not too sure but by the general look of it, it seems to be the leaf beetle Chrysomela populi.
This is mostly from the idea that they are very common in Europe and the distribution of the spots on its body. If nothing else it is most likely from the Chrysomela genus.
Here is a link to a description of the species

For species ID questions, we ask that answerers provide a reference for their ID, and preferably a description of what led them to that ID or defining characteristics (such as things that allow you to exclude other possibilities).


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How much does dry moose scat weigh and vary in dry weight?

I am in the early stages of designing a study involving a moose (Alces alces) population.

One of the important objects of measure are their moose pellets, which I want to take a battery of measurements on including their dry weight.

I recently picked one up off the forest floor and subjectively it felt quite light.

Originally I had intended to buy a ~$100.00 weigh scale with 0.01g precision.

If the pellets are too light to be detected by the balance, that is a obviously problem.

But I am also interested in variation, so it would also be a problem if most of the variation among pellet weights occurs below the precision of the balance.

Has anyone published statistics on the weight and variation in weight of moose pellets?

Why individual pellets rather than pellet groups? But 0.01 g is quite small...

@BryanKrause If I measure aggregates of pellets then I cannot study their individual variation. Why I care about their individual variation is particular to the study questions.

@BryanKrause The precision of 0.01 g is quite small, subjectively. I want to know ahead of time, if possible, that I have sufficient precision.

MacCracken and Van Ballenberge, 1987 report that pellet size varies a bit by sex and age; male pellet dry weight (all measurements are dry) averaged 1.9 grams (range 0.9-2.9), female 1.3 g (range 0.6-2.0), yearling 0.8 g (range 0.6-1.7).
Unfortunately they do not report SDs but instead SEs of 0.01 for every group; we can try to reverse engineer the SD but with one significant digit there isn't much precision to work from. SE = SD/sqrt(N), their N were 34/36 for the male/female adult groups, so let's just base it on 35...
0.005 * sqrt(35) = 0.030 g
0.015 * sqrt(35) = 0.089 g
The SD could be somewhere in that general range, but based on the range I suspect it may actually be much higher, and that they used a larger N for calculating the SE (perhaps many pellets per group). The other possibility is that the range is mostly due to outliers. As a rough estimate from the ranges, you could guess the SD is around 1/6th of the range (if normally distributed, 99.7% is within +/- 3 SD), with ranges around 2.0 to 1.4, that would give you a SD around 0.2 to 0.3 grams.
They used a scale with 0.01 g precision; seems sufficient for this work. Dry weight is certainly the standard for this sort of thing, so you may want to look at equipment for drying as well.

MacCracken, J. G., & Van Ballenberge, V. (1987). Age-and sex-related differences in fecal pellet dimensions of moose. The Journal of wildlife management, 360-364.

(+1) Thank you for (1) finding a citation and (2) clarifying whether it was dry/wet weight.

Nice use of the definition of standard error of the mean, btw.

@AgnesianOperator I think likely my original estimates for SD based on SE are very low; I suspect they put in measurements for many pellets per group and calculated their SE across all of them, so the N=35 is wrong. I think my new estimate based on the ranges will be closer to what you actually observe.

Thanks, I appreciate that update to your answer. Regrettably, I can only upvote your answer once. I will read the study to understand their statistics in detail. On the face of it I am thinking of using a random intercept model to allow different piles of scat to be from possibly-different individuals whose age/sex might differ.

@AgnesianOperator Yes, I would certainly suggest mixed effects models for something like that. I don't think you'll find much use in the stats for this particular paper, as although they are doing some interesting stuff for 1987, statistics has come a long way, particularly in how biologists use statistics.


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What is the average rate of red blood cell production in humans?
I was doing my biology research of the day and I came across the following question: How fast are red blood cells produced and, how do I calculate this?
I am currently using the following:
Calculated estimated blood loss (cEBL) = (Predelivery HCT- postdelivery HCT) / predelivery HCT.

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

One thing I suggest you look up is how RBC form (if that isn't what you are asking about, please remove that tag) — that should help you to phrase your question more clearly. I'm going to copy edit your post ("blood reproduction speed" and "blood reproduction rate" aren't clear and "how fast is the ... rate" is poor writing) to start to improve its clarity, but please check carefully to ensure I don't inadvertently change your intended meaning.

In addition, your equation is about blood loss not RBC generation — please add why you think that is relevant or edit the question so that that equation makes sense. Also note that all abbreviations must be defined before use — you should not assume we all know what HCT means.


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In life history theory, does trade-off between reproduction and growth and maintenance imply longer health span for a sexually inactive organism?
The life history theory suggests that there exists trade-off between life processes like reproduction and growth and maintenance of the organism.

One significant trade off is between somatic effort (towards growth and maintenance of the body) and reproductive effort (towards producing offspring).

Quoting from wikipedia page Cost of reproduction hypothesis

The traditional view of the cost of reproduction says that a cost is caused by differential allocation of limited internal resources. (...) Reduced immune function and a decreased defense against environment stress are also important factors in the cost of reproduction.

It has been suggested that this kind of trade-off in resource allocation between different life processes has plasticity, that is, the resource allocation between different life processes in a given individual is dependent upon external factors like the absolute amount of nutrient input.

Reduced nutrient availability can substantially magnify, while increased nutrient availability can diminish or obviate an apparent trade-off. (...) For example, in laboratory and field experiments on bivalves, cladoc-
erans, insects, and mammals, allocation to maintenance or storage was found to
take precedence over allocation to reproduction under nutrient-poor or stressful
conditions

So, my question is, what does all this mean? Could sexually inactive life for an organism, for whatever reason, actually correspond to longer health span? (Better immunity, growth and maintenance of the body, etc.)
I know that the rate of deterioration of health with age is hugely determined by the rigid parameters corresponding to the specie the organism belongs to. But for a given organism, the various life processes like reproduction, repair and maintenance, and immune system are fighting for the limited resources available, and it appears that there is some amount of plasticity in how much resource will be allocated to which life process. In other words, the rate of deterioration of health of an individual appears not to be completely written on stone.

I am not sure if life span could be extended in such circumstances but the probability of survival could probably increase, given the high metabolic costs of growing and raising offspring. I work mainly with marine mammals and there are multiple studies showing that the body condition of females following births is much lower (as to be expected), and thus health indices are worse. Therefore, these animals are at greater risk of being severely affected by infections or variable densities of available prey, showing to an extent the high costs of reproduction.
Another thing to consider is the level of stress that any member of the population is exposed to. Whether an individual is sexually active or not, various stressors (i.e., limited prey, pollutants that compromise the immune system) can limit their life span and increase mortality risks. For example, there are various studies on the effects of persistent organic pollutants such as PCBs (i.e., https://pubmed.ncbi.nlm.nih.gov/29096314/) that can be passed down to calves and persist in the body for long periods of time. These substances can compromise the immune system, thus putting animals with high concentrations of these pollutants in their bodies at greater risk irrespective of whether they become sexually active or not. Therefore, there might be a higher likelihood that an animal can live (potentially) longer as a result of the lack of investment in reproductive activities, because the resources can indeed be used for growth and body maintenance, but there are stressors that can limit survival rates before the animals are even born.
Here is a study that is not directly related to the cost of reproduction, but investigated the plasticity of immune investment in individuals exposed to human-generated stressors and limited food availability: https://journals.plos.org/plosone/article?id=10.1371/journal.pone.0067132
I hope this answers your question and if you are interested in references to papers that were briefly mentioned but not specifically provided, I am happy to help!

You bring up an important factor with the human generated chemicals likely to contaminate much of what's currently likely to be observed.

I wonder if this also relates to matriarchal societies needing their matriarchs to survive longer (to lead, pass on knowledge, etc) and menopause/cessation of female fertility?

Also keep in mind that females can offload contaminants to offspring via gestation and lactation, therefore reducing the contaminant load of the mother. But whether or not reduced contaminant loads offset the energetic demands of motherhood is another question...


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Hierarchial and shotgun sequencing vs Massively Parallel Sequencing
Why is the cloning step unnecessary prior to sequencing with massively parallel sequencing (MPS; Illumina or similar technologies), when it is necessary for hierarchical and shotgun sequencing?
I know that the cloning step is helpful for introducing universal primer binding sites for sequencing. So is it because we are using probes with adaptors on them for MPS that we don't need to clone?

Can you clarify what MPS is or link to some information that describes in more detail what you mean? I can imagine that "MPS" is "massively parallel sequencing" but that acronym is not standard terminology, as far as I know.

@MaximilianPress Oh sorry, yes I mean massively parallel sequencing.

The construction of sequencing libraries for Illumina, PacBio, ONT, or similar  instruments is a large field of methodology that encompasses a variety of different techniques. For background on Illumina libraries specifically you can see here.
In some cases, notably Illumina, indeed the library construction involves sequencing adapters either being ligated or added by PCR. There are many strategies here and I don't claim exhaustive knowledge of all of them, I would suggest doing more research here if you are interested.
For other cases, sequencing occurs by alternative mechanisms. I suggest consulting overviews of PacBio and nanopore (ONT) sequencing to get ideas of how different methods work for the same basic task of shotgun sequencing of DNA without a cloning step.


text:
ant ID montreal, qc canada
I'd like to know what type of ants this is. These appeared in my basement recently.
I am thinking those are called "pavement ants" after comparing online. Please help me identify them.
( Sorry for the blurry image quality, these are quite small and are scared of my camera )


text:
Why can $\alpha$- and $\beta$-glucose molecules not link together?
My textbook states that for two glucose molecues to link together, they must be of the same form, i.e. they must both be either $\alpha$-glucose or $\beta$-glucose molecules.
Why is that the case?

What precisely does your textbook say? Can you quote and cite? What is your thinking on the matter/best guess from what you know?

You tag your question organic chemistry, and perhaps you are thinking of it from that angle. Think of it as biochemistry.

@BryanKrause my textbook says: "Although some chains of sugars store energy, others serve as structural material for cells. For two glucose molecules to link together, the glucose subunits must be of the same form. Cellulose is a polymer of $\beta$-glucose." I don't see any reason why an $\alpha$- and a $\beta$-glucose molecule shouldn't be able to link together, chemically. The only other idea I have is that maybe there are no cells/enzymes that can facilitate a dehydration reaction between the two - which would fit David's hint.

It would indeed fit because enzymes only evolve and “stick” if they catalyse a reaction of use to an organism. Your textbook should have made that clear.


text:
Entry of particulate pollutants into the nasal cavity
NCERT Chemistry of Grade XII (India) writes

Particulate pollutants bigger than 5 microns are likely to lodge into the nasal passage, whereas particles about 10 microns enter the lungs easily.

I'm primarily a Mathematics student, and my experience in Biology has been fairly limited to high school Biology, so excuse any factual errors that I make during my argument.
From what I know of the respiratory system, the trachea divides further and further getting narrower (into Bronchi, Bronchioles), so how does this happen that particles smaller are getting lodged into the nasal passage, whereas larger particles are getting into the lungs? Shouldn't it have been the other way around?

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

Yes indeed, your quote has things backwards. Here is an image and a link to further information:
ref

If you follow the dotted line with your eyes - - - - - through the 1Micron grading... On the right side, the majority is absorbed by the head. On the left side, it's mostly the lungs.

other references


text:
How to mark nest area in double bridge experiment?

In my setup, I am lacking a nest. I'd need some way of telling the ants to bring food back to a first container, instead of just accumulating at the food source.

Presumably you have a queen, or at least some larvae?

Ah yes that might work, or maybe I could put one end of the setup close to a nest outside

Could you please explain what this is all about and what your question about biology is.


text:
Why do some people accumulate more diglycerides in their muscle cells?
The scientist Gerald Shulman has experimentally found that young lean adults in their early twenties that are children to people with type 2 diabetes often show muscle insulin resistance. He found that diglycerides within the muscle cells made it harder for the insulin activated "glucose transporter 4" to attach to the cell membrane.This causes these people to have raised insulin levels in order to get enough receptors to get glucose into the cells.
Also he has found that young adults of the same age and BMI that are not "sedentary" but more physically active do not have insulin resistance in the muscles.
Shulman believes that muscle insulin resistance due to diglycerides in muscle cells is what is starting a series of events that eventually can result in insulin resistance of the liver and diabetes. He has more then 450 published papers on related subjects and I can not really find "the one paper" where he claims to experimentally have verified his assertions.
This link is from a presentation from when he won the Banting Medal:
https://professional.diabetes.org/webcast/banting-medal-scientific-achievement%E2%80%94mechanisms-insulin-resistance%E2%80%94implications-obesity
There seems to be different theories as to why some people tend to accumulate more diglycerides in their muscle cells.
Question: What are the major reasons some people tend to accumulate more diglycerides in their muscle cells than others?

Remember the one question per post rule we have here? Asking so many in one post is liable to result in less than full coverage of the topic.

Further to @ARogueAnt., Asking so many questions is likely to have your question closed.  This is Q&A site, not a journal club or a discussion site. It might be worth your while looking at the Help on Asking Questions.

Thank you for focusing down to a single question, but I think you would get a better response if you supplemented the youtube links with references that supported the assertions you make. That would also demonstrate that you had done the expected research. ——— This is a good example of how to format references. Also note that tags should be created sparingly, should never be redundant, and are most useful when they apply to multiple questions.

Having already made a criticism of your question I am reluctant to make another, but please accept it as having constructive intent. Professional scientists have an aversion to individual scientists publicising their views on videos. That is not how science is done. Like @tyresome, I read peer-reviewed papers and look at the evidence, not the person. A search for "diglycerides and insulin resistance" brought up a number of papers on this topic, implying that the correlation is well known. I suggest you read these yourself to see if there is any work on the cause of increased DGs.

@david I can see that this is a very active field of research, some clinical trials of diabetes-oriented medicine regulating lipid metabolism seem to be coming up. Some researcher thinks that fructose consumption via some pathways destroys lipid metabolism and thus causes insulin resistance. I am trying to lift the knowledge in this area (which do not seem to be widely spread) to a wider audience. For that I need answers from people with higher ranking than me on this site.

Clinical trials don't give information about mechanism. Neither do researchers' thoughts. If diglyceride synthesis increases as a response to some change, the mechanism by which this occurs can only be elucidated at the cellular and subcellular level. There are not too many biochemist frequent this site, and this is a rather specialized area. Sometimes I would try to research the literature out of interest, but I'm not inclined to do so as I'm busy and suspect that it may be a wild goose chase. You might try on SE Medical Sciences if you get nothing here, despite your bounty.


text:
Difference between in cis/in trans and in phase/out of phase
Something I've never been quite clear on is the difference between in cis/in trans and in phase/out of phase, as used for the genetics of diploid organisms.
My understanding is that the in cis/in trans terminology is used specifically when referring to the configuration of recessive and dominant alleles for a pair of genes, while in phase/out of phase is used when referring to a pair of individual variants, such as SNPs or indels, however it would be great to get an authoritative clarification on the issue.

Like all words, the meaning of terms like these strongly depends on context — quotes (with appropriate citations) would probably be your best bet for getting a clear answer, but at very least you need to provide more information than "in diploid organisms".

Posts must be self-contained — if I need to search on youtube to understand your question it is not finished! 2) You are expected to demonstrate that you have done the expected prior research — that means consulting textbooks and other reliable sources before posting. Youtube videos are of highly variable quality and I don't have time to vet them. Please [edit] your question and tell us where you've looked for answers (not "youtube"), what you do know about the topic, and where exactly you still have questions. Please take the [tour] and consult the [help] starting with [ask] for details.

Unless there are other meanings of the terms I'm not aware of, they are referring to slightly different things.
Haplotype 'phase' refers to whether alleles are found on the same chromosome (paternal or maternal). Hence, 'phasing' refers to the statistical / read-based process of determining which chromosome a genetic variant is found on. 'Unphased' data, typically produced by genotyping arrays or sequencing technology (e.g. short Illumina reads), doesn't contain information about which chromosome a variant is on.
In some contexts, cis/trans terminology refers to whether a locus (e.g. a eQTL/pQTL) is found near the gene it corresponds to. For instance, say you run a GWAS on protein levels and find a significant hit for a locus controlling the level of the protein X. If this locus is within some kind of threshold distance of the gene encoding protein X, perhaps 2Mb, then it is known as a 'cis' ('on the same side as'). Otherwise, if the locus is found distal to gene encoding protein X, it is known as a 'trans' locus. Similarly, 'cis' regulatory elements are regions of non-coding DNA which regulate the transcription of nearby genes.
For example, from this paper:

In the cis-eQTL analysis, we evaluated the associations between lncRNA
expression levels and the genotype at a given SNP locus located within
1 Mb upstream and 1 Mb downstream of lncRNA. trans-eQTLs are defined
as associations beyond the 2-Mbp interval.

I took a closer look a number of Wikipedia articles, including those you linked to. The Latin root of cis/trans means this side of/the other side of, which calls to mind pairs of chromosomal homologs, however modern usage of the terminology appears to mean something more like same molecule/different molecule. Whether or not acting "in cis" requires that the action is in some way facilitated by the genetic elements all belonging to the same DNA molecule is left unclear, and "in trans" is still sometimes used for distant but still chromosomally co-located genetic elements.

Can you provide a citation for the use of cis/trans that you give — you seem to be saying it is primarily about proximity, which conflicts with the usages I've encountered in biology (and chemistry) ...

@tyersome could you be more specific about what use I refer to that you're interested in more information about? If you mean the use stated in my original post, then just go on YouTube and type in "cis trans genes", the top hits will all be videos explicating the so called cis/trans configurations of recessive and dominant alleles.

@Thoth given there seems to be different definitions of cis/trans, you should really give the context with which you are asking this question.

@user438383 in the context of genomics/genetics.

Ok well my answer is in the context of genomics.


text:
biofilm model scales
I have a question about the meaning of biofilm modelling scales:
Are they microscopic, mesoscopic and macroscopic.
Microscopic means individual bacteria
macroscopic means large number of concentration
and mesoscopic means the structure of biofilm.
If there is any mistake please correct me.
Thank you

I'd have thought it rather depends on the context in which the phrase is used. Could you elaborate? Is it a large bioreactor or a petri dish?

Thank you for your response. Actually, I am using mathematical models to model bacterial biofilm. However, I am required to write about biofilm modelling scales and to be honest I ve read several papers and I am not sure if my conclusion were correct.

Just like in math, in biology you have to define your terms. "Modeling scales" are whatever someone defines them to be, within the context their definition is used. Two papers can use the same word to mean something different if they want. For your writing, you'll have to cite the actual literature relevant to your field, not an answer on StackExchange.

I am not going to cite StackExchange. Just I wanted to clarify my understanding to the concept.


text:
How could microplastics accumulate in the bodies of marine mammals?
I have read several literature reviews and studies on the effects of microplastic particles on fish and invertebrates (one example includes the review by Franzellitti et al. (2019)) and there are multiple instances where these particles where found in the nervous system, lymph, and other organs following controlled exposures.
There are also multiple studies that looked at microplastics in marine mammals, but usually they focus on the contents of the gastro-intestinal tract. My question is: how these particles are likely to accumulate in the bodies marine mammals compared to smaller organisms - could the size of the blood vessels of larger whales influence the uptake of microplastics? Some researchers recommend studying this by using marine mammal cell cultures, but I am wondering how well the results of such studies can reproduce what might be happening at a full-body scale. (I study mainly ecology and the behaviour of marine mammals, so my knowledge of differences between various organisms at a cellular level is limited).
Thank you!

Microplastics are bioavailable to marine organisms, through direct ingestion and/or via trophic transfer. Most studies of microplastics in marine mammals only analysed microplastics in the digestive tract and scats (see Zantis et al. 2021 for a review) and thus, at this stage, there is no evidence that microplastics can cross the intestine barrier and/or enter the blood stream and accumulate in different organs in marine mammals. However, in some marine vertebrates (fish), microplastics have been found in other organs (e.g., brain, liver).The small size of microplastic might allow them to pass through the intestinal epithelium barrier through cell internalization and possible subsequent translocation. However, information concerning the extent of ingestion, accumulation, translocation into organs and possible pathways of transition into cells is still unclear (Wright et al. 2013).
Interestingly, it has been very recently demonstrated that microplastics are found in human bloodstream (Leslie et al. 2022). It is scientifically plausible that plastic particles may be transported to organs via the bloodstream. Regarding your question, "how these particles are likely to accumulate in the bodies marine mammals compared to smaller organisms", it's a difficult one. You need to consider the exposure route (direct ingestion and/or via trophic transfer) and the particle size (and the bioaccumulation and biomagnification processes).


text:
What is meant by “global RNA polymerase occupancy”?
I read the following in a paper

It provides base-pair resolution and strand-specific information of **global RNA polymerase** occupancy.
CDK13 cooperates with CDK12 to control global RNA polymerase II processivity

What does this mean?

Please provide a reference to the article/textbook you are citing.

"Global" tends to be used in contrast to "local". I think you need to read the paper to find out more specifically.

it looks like you are posting quotes without acknowledging them as as such or noting where they come from. It would help for you to clearly restate a more expansive version of your question outside of the title for us to be able to help more effectively.

In context of the PRO-seq method, "global" can be substituted for "genome-wide". Note that use of the term global is a call-back to the methodological predecessor of PRO-seq, which was called global run-on sequencing (GRO-seq).


text:
Confusion regarding the meaning of the coding strand of DNA

I am having problems getting the ‘correct’ answer for the above question.
I am assuming that mRNA uses the template strand (not the coding strand) for protein synthesis. So:
         ---------[DNA]  -  [mRNA]  - [Amino Acid]

Original A T A -> U A U -> tyrosine
Mutated  T C T -> A G A -> Arginine
What is my mistake?

Welcome to the site. Please visit the [help] and take a [tour]. One of the requirements for questions on this site is that they demonstrate that they have done the research to look for the answer themselves. Think about what the question is asking and what the word coding means...

Look up "coding strand" in wikipedia.

The problem here is the misleading nature of the term “coding strand”.
I have previously posted on this topic making the point that mRNA — which is what the genetic code relates to — should be the basis of nomenclature, and that the best and least ambiguous terminology is sense (the way the code is read from the mRNA) and antisense (its reverse complement). To quote myself:

The idea seems to me clear that when you read the string of codons
that encode the amino acid sequence from this strand they ‘make
sense’. (Anti-sense is used in preference to non-sense, as ‘nonsense’
was the term used historically for mutations that converted amino acid
codons into stop codons.) It can also be extended to non-protein
coding genes (e.g. for tRNA), where ‘sense’ correlates with the
sequence of the gene product.

Thus, if you consider the two strands of DNA, the one that is transcribed by RNA polymerase into DNA is the anti-sense strand, and the other (which is the one humans may read after a DNA sequencing experiment) is the sense strand.
My main objection to the use of coding and non-coding for DNA strands (apart from logic — how does one code and the other not?) is ambiguity:

As the anti-sense strand is the template for RNA polymerase, one might
make the mental association between this and ‘coding’, whereas it is
the sense strand that is meant [by coding] in this (admittedly common) usage.

So in my opinion the question is to blame, although this is very common. But, to answer the question, one presumably should assume that the strand with the asterisk is the sense strand, and any changes in it are reflected directly (apart from T to U) in the mRNA.
Footnote
It is ironic that the genetic code provided in this question is unlikely to be correct, although the organism is not specified. For example, in vertebrates the following applies:
 AGA    Stop (not Arg)
 AGG    Stop (not Arg)
 AUA    Met  (not Ile)
 UGA    Trp  (not Stop)

[https://www.ncbi.nlm.nih.gov/Taxonomy/Utils/wprintgc.cgi]


text:
Is it scientifically sound to pool repeated measurements?
I am measuring specific phenolic compounds in leaves of A. thaliana. I have many different varieties, and different treatments. Initially I was measuring one leaf per plant for three plants for each treatment, cultivar combination (left in example image). Taking leaf punches is the fast part, actually processing the samples is very time consuming and rate limiting. To increase my accuracy and throughput, it would be nice if I could sample more than 3 plants per treatment,variety combination! In this case the standard deviation should represent the differences inbetween samples of the same treatment, cultivar combination.
Therefore, I am now opting to mix the three samples from the three different plants from the same treatment,cultivar combination and repeat this two or three times (right in example image). This way I will lose measurements of single plants, but I should still be measuring the average of my treatment, cultivar combination. By doing measurements this way, it also means I can pool more than 3 plants, for example 6, in a single sample, while keeping the same amount of processing work afterwards. In this case the standard deviation should represent the accuracy of my laboratory practices, but the mean is actually closer to the true mean.
My question is if this is scientifically sound to do?

I’m voting to close this question because it is much more appropriate for CrossValidated

Why are you pooling the samples multiple times? Are you planning on doing different analyses on the different samples?

After asking why you don't think circumstances alter cases, I'll go with Ben Bolker.

I am pooling the samples multiple times to get some sort of a standard deviation, as said it should represent the difference between pooled samples and/or accuracy of my assay

Your intuition is largely correct here.
Bryan Krause's comments focus on where your answer is wrong.
Some of his statements are true, others are speculative.
His answer, combined with your intuition, could improve your measurements.
I've shown a pooling scheme below.
It should address the key ideas from both your question and Bryan's reply.
The two key ideas here are (i) that pooling can improve estimates and (ii) that repeated measurements can give you insight into sampling variation.
Each observation you make will show a different amount of the phenolic compound.
Some of this is due to actual differences in the amount of the phenolic compound.
These differences can exist between fields of plants, between leaves on a plant, between punches from a single leaf.
The list goes on.
Furthermore, each measurement of a single sample produces a different reading.
This is a lot to account for.
If you would like to use statistics, then you should make it clear how your sample relates to the population you are interested in.
You mentioned that taking leaf punches is relatively easy.
Pooling leaf samples uses that to your advantage.
Let's say you will only be making three measurements for cultivar A and three measurements for cultivar B.
For the first measurement you could grow 9 plants of cultivar A, and take 4 punches from each of those plants.
You mix these punches together and measure the abundance of your phenolic compound.
For the second measurement on cultivar A, you could repeat this entire process using different plants.
The difference in these two measurements reflects sampling variation.
This is due to all the sources mentioned above, and others.
Decreasing the number of plants you pool doesn't solve the problem of sampling variation.
From a statistical standpoint, pooling can allow us to average.
Lets say punches $i = 1, 2, \ldots, n$ are taken from cultivar A.
They are prepared as equal volume samples.
The corresponding concentration of the phenolic compound in each sample is $A_i$.
Mixing these solutions results in a sample with concentration $\bar A_n$.
The formula for this concentration can be obtained from the concentration of each $A_i$ as follows
$$
\bar A_n = \frac{1}{n} \sum_{i = 1}^n A_i
$$
These concentrations would be different if we repeated the experiment with a new sample of punches.
To capture this, we can treat them as random variables.
We would like to show that pooling decreases the variance of $\bar A_n$.
To see when this can happen note
$$
\text{Var}\left[\frac{1}{n} \sum_i A_i\right]
= \frac{1}{n} \left(\frac{1}{n} \sum_i \text{Var}[A_i]\right) + \frac{2}{n^2} \sum_{i < j} \text{Cov}(A_i, A_j)
$$
The leading factor of $\frac{1}{n}$ is a key term here.
It has the potential to make the variance small when the number of punches $n$ is large.
This highlights the benefit of pooling, and is consistent with your suggestion.
A pooling strategy. Each tube measures a mixture created using multiple punches. Each punches is measured in a single tube.

Thank you, I think I get the core message in your story :). I.e. For a single treatment/genotype combination, I could do three measurements, every measurement containing pooled samples of multiple unique plants!

If you use the configuration on the left, your variance across tubes reflects variance between individuals.
If you use the configuration on the right, your variance across tubes reflects primarily variance in your assay (and perhaps variance within a leaf a bit).
Usually, you do an experiment like this because you'd like to be able to extrapolate your results to a larger population. To do that validly, you need to be able to estimate the variance in the population from your sample: the variance between individuals. Your configuration on the right will help you tell whether your samples are internally different, but they don't tell you much about what you can expect outside your sample. Importantly, one outlier plant would contaminate all the samples it contributes to.
Note that unless your effect sizes are expected to be very large, 3 individuals per group is likely an underpowered sample, meaning you do not have enough observations to detect a true difference in your groups. You cannot get around this by taking more samples from the same individuals, you need more individuals.
There are some cases where you might use an approach like on the right to test your assay, though not to draw conclusions about differences in treatments.
The language we use in statistics to describe these scenarios is independence. For, say, an unpaired t-test to be valid, the samples must be independent, both within and between groups (same is true for ANOVA equivalents with multiple groups). In the right scenario, your samples are not independent: every sample that includes part of "leaf 1" is going to have a relationship with every other sample that includes part of "leaf 1". If you treat those samples as if they are independent, you are breaking the assumption of your hypothesis tests and cannot rely on the results when comparing groups.

For a bit of an analogy that moves away from the more complicated biological assay part, let's say you want to know whether Group A is heavier than Group B. The procedure on the left is like weighing each member of Group A, and comparing these weights to members of Group B.
The procedure on the right is like having all members of Group A stand on a scale, then get off, then all stand on a scale again, then get off, etc. You'll have the best measurement you can get of how much the specific sample of Group A weighs across different measurements on your scale, but you won't be able to use your sample of Group A to estimate how much variation there is in Population A: your variation will just tell you how reliable your scale is. This is clearly a very very wrong procedure that is equivalent to the approach as you are proposing on the right-hand side of your question. Do not take this approach, you will waste your time and your results will be incorrect; if you hide the fact that you've taken this approach your work will be fraudulent.

Thank you. You state a wrong procedure, but you also say it gives the same result. In my case I can have a much higher and more accurate throughput with the wrong procedure. Therefore, from a non-scientific perspective, this method could be preferred?

@Rivered It definitely does not give the same result. Do not use the procedure depicted on the right to compare your groups, you will have wasted all of your time or committed outright fraud if you do. Not sure where you got the idea that it's the same result, but if you can quote from my answer what made you think that I will change it ASAP.

I toke the remark from your statement "it's the same design as". In any case I understand your answer, and statistics are useless without estimations of variation. The whole point of my questions was the throughput, with the "wrong" procedure I will be able to measure 10-fold more samples, and although missing variance, the group mean will be much more accurate due to the high sample number.

@Rivered In the analogy section I am describing a procedure that is very wrong, yet is equivalent to the design you propose on the right-hand side of your question. Do not do this procedure. And no, the mean measured the way you propose will not be any more accurate as a population estimate. "Less variance across samples" does not mean your mean is more accurate, it just means you estimated the variance wrong.

I am not saying my mean is more accurate because of the lower variance! I say my mean is more accurate due to the ability to measure much more samples in a pooled way. To put it in terms of your analogy, I can place 100 people on a single scale, whereas time constraint wise I could only weight 3 people seperately. The mean of 100 people will be much more close to the true value, compared to the mean of just 3 people?

@Rivered That is not what your question depicts. If you have a new question, probably stats.SE is a better place to ask.


text:
Is this bark beetle going to be a problem?
This little (around 3mm in length) beetle is one of many that infested a gingerbread heart in our house. It seems to be some kind of bark beetle. My question is what exact species it is likely to be and whether it is a danger to our wooden floor, furniture, maybe even roof. We live in Germany.

Stegobium paniceum. (Biscuit beetle, bread beetle, drugstore beetle).

Copyright Siga, via Wikipedia, CC BY-SA 3.0 2022.

Copyright marked, via entnemdept.ufl.edu, fair usage, 2022.
Characteristics:

Adult can be up to 3.5 mm in length.
Worldwide distribution though it is more common in warmer climates.
Notched antennae (like the teeth of a saw).
Diet consisting of a diverse range of dried foods and spices, as well as hair, leather, books, birdseed, dried pet-food, and museum specimens. The drugstore beetle is also known as the biscuit or bread beetle since it can live on biscuit or bread crumbs.
Larval form is a small, white grub. Period in this phase can be several months. Larvae do most of the damage this creature causes (great economic harm).
Likely to eat through foil-sealed food packaging.
Lives in obligatory symbiosis with a yeast fungus.

Pest control measures:

Thorough cleaning.
Destroy infested material.
Freezing below -20 Celsius (-4 Fahrenheit).
Diatomaceous earth sprinkled in corners may help.

Resembles, but not to be confused with a related species:
Anobium punctatum. (Common furniture beetle, woodworm).

This seems almost certainly correct. Not sure this is great news, but at least it's not going to bring down the house.

I taught Latin grades 3 -12. That was the magistra in me bursting out, sorry. :-/

Not at all. It's just that no-one taught us about pre-classical Latin. Never even thought to look anything from that period up before. I'm all too glad to learn. @anongoodnurse

I really like the etymology of words (English, Latin, French, Norse, whichever.) It never occurred to me to stop at some arbitrary line.

For me, it's not so much that I stopped at a line, just that I've been given snapshots of languages at school, and I've continued taking them, increasing the range of languages as I go. Hadn't gotten around to looking at what came before Catullus, Cisero etc., except when looking at Ancient Greek words that get kicked-up in English from time to time. My paternal aunt is the real linguist of the family with her Russian, Spanish, Italian, Bulgarian and a heavy dose of comparative linguistics. Weird to think that she never used it in any of her jobs. @anongoodnurse

Wow!!! Kudos to her, seriously! French was my first language, so the Romance languages are easy (Well, not Portuguese) and Latin, of course, informs those. Kudos to you, too, for following Greek. Once the letters change shape, I wimp out. :( Only used French twice in my decades as a doc. Spanish? Nearly every day.


text:
Is Zayed et al.’s revision of Orthornavirae robust?
Zayed et al. (April 7, 2022) in Science “Cryptic and abundant marine viruses at the evolutionary origins of Earth’s RNA virome” report several major hitherto-unknown Ribovirial phyla:

The two most sampled, Taraviricota and Arctiviricota, had the lowest normalized abundances.
To mitigate issues of subjectivity, their RdRp-based phylogenetic analysis was developed to recapitulate the established ICTV-accepted taxonomy (97% agreement).  Their new tree shows a very different early evolutionary history, for example splitting Duplornaviricota into 3 distant phyla:

Are these findings robust?  The unknown megataxa and the revision of Orthornavirae?

Do you really think that there is someone on this list who has read and is better able to assess this paper than the referees assigned to it by Science? I presume you feel it important — I have no idea — but if it is I’m afraid you will have to wait either for direct responses to the journal, or future publications citing it. This is a question and answer site  about biology, not a journal club.

Can you identify the evidence supporting alternate tree topologies? If you show that evidence here we might be able to judge it.

Paper certainly looks robust - you would need to dig out the original data and analyse for yourself if you wished to compare. You can find the annotated data here: https://datacommons.cyverse.org/browse/iplant/home/shared/iVirus/ZayedWainainaDominguez-Huerta_RNAevolution_Dec2021

@David Sometimes in phylogenetics the reviewers of science are just a joke (see the Omicron evolved in west africa retracted paper where the non-omicron raw reads were just contaminated with later omicron strains)

The main difficulty here is that it is risky to feed mafft directly with so distant RdRp, the resulting alignment will be a mess for sure. You can try running a MCMC optimizing both the tree and the alignment (after subsampling the dataset) and see if the deep branches topology stays comparable.