text:
How do i create sustainable readily available stock cultures to act as Gram stain controls?
My investigation on the matter
Ref #1

Staining known Gram-positive and Gram-negative organisms on either
side of your unknown organism act as positive controls for your
technique.
(Microbiology Laboratory Theory and Application (Michael J. Leboffe, Burton E. Pierce)

Ref #2

Similarly, cultures should undergo evaluation while they are still
fresh. Old cultures tend
to lose the peptidoglycan cell walls, which predisposes gram-positive
cells to be gram-negative or gram variable.
(Tripathi N, Sapra A. Gram
Staining. 2023 Aug 14. In: StatPearls [Internet]. Treasure Island
(FL): StatPearls Publishing; 2023 Jan–. PMID: 32965827.)

Ref #3

Refrigeration impedes further growth of the bacteria, so you will have
"fresh" cultures to work with during the next laboratory session.
(Laboratory Exercises in Microbiology By Robert A. Pollack, Lorraine Findlay, Walter Mondschein, R. Ronald Modesto)

Ref #4

Remember the following points concerning your [refrigerated or frozen] slant or agar stab stock
cultures:
-Do not use stored stock culture for making slides or routine inoculations.
(Laboratory Practices in Microbiology By Osman Erkmen)

Question
How do I create a sustainable readily available pair of stock cultures to act as Gram stain controls (as Ref #1), in the least time-consuming and tedious way?
From Ref #2 and #3, I can prepare two slant cultures, one is of a Gram-positive bacteria and the other is of a Gram-negative bacteria, then store them in the refrigerator or freezer. Then I can retrieve them daily and use them. After that, they are returned to storage.
However, Ref #4 argues against using refrigerated/frozen cultures for making slides.

Just spitballing, but what if you made a stack of slides and heat fixed smears of your controls onto each slide, so that you could just grab one out of the slide box and have your controls already fixed to the slide. It would probably take some verification testing against fresh controls to determine the quality/stability, but I know I have used commercially available QC slides like this in the past.

It's not about the mean, but the end. Anything that would substitute as a Gram stain control would do. So this is an answer.

Aside from MikeyC's commented suggestion, which I think is a good idea and they should write it up as an answer.
In my experience, most species of bacteria you might use as controls for Gram staining will easily last 2-4 weeks in the fridge on a plate and still be usable as controls.  These will be genera like Staphylococcus and Escherichia, well known, readily available and well characterized.
For fairly minimal effort, one can simply take a plate of the organism wanted and re-streak/sub-culture once a week or once every two weeks. The streaking itself should only take you a minute or two, adding in time for labelling of plates and general fluffing around in the lab (opening incubator, placing plate in incubator etc.), maybe a maximum of 5-10 min per plate. I know that with pre-labelled plates and disposable loops, I can sub ~100 plates in about an hour.
For longer term storage of species, it is best to keep a glycerol stock. These are stored at -80 C (ultracold freezer) and can be used indefinitely. If you wanted to work from glycerol stock (to be sure of getting the right species), add maybe another 5 min to look up location in the freezer and get them out for streaking. There is no need to thaw a glycerol stock (in fact it is detrimental to thaw them) - just use a cooled flamed loop, or a clean disposable loop to scrape a tiny amount of the stock from the top of the tube and streak this out.


text:
Does respiration require energy?
This question in my class 10 test and my answer was "since in respiration glucose is broken down into pyruvate, thus chemical(enzymes) energy must be required." I also thought breathing is a part of respiration, this requires the lungs to be moved- thus energy required.
But my teacher says otherwise. She said I am wrong and her explanation mainly circulated around how respiration is exothermic etc...

Cellular respiration.

Sorry, couldn't grasp what the site said.

The first step in respiration is the breakdown of glucose into pyruvate. This reaction occurs in cytoplasm and is called glycolysis. Glycolysis occurs in a series of reactions. In fact, the first half of glycolysis is referred to as the 'energy requiring phase' which uses 2 ATP.
https://www.ncbi.nlm.nih.gov/books/NBK482303/
In this article, you can get its reference.

The first phase is the "investment" phase due to its usage of two ATP molecules, and the second is the "payoff" phase. These reactions are all catalyzed by their own enzyme, with phosphofructokinase being the most essential for regulation as it controls the speed of glycolysis.

But, as others have pointed out, Respiration has no net requirement of ATP. Even, no. of ATP molecules produced are based on net production. So, it's appropriate not to consider respiration requires energy.

But glycolysis overall has no net requirement for ATP, and the way that the question was posed suggests that the overall process was what was meant. Certainly the argument of the poster were not the "phase I, phase II" of glycolysis, and the illogicality of his argument and his confusion with the role of enzymes does not suggest that he is even aware of this. You do him no service by suggesting his answer is in any way correct.

@David The poster did try to explain that phenomenon as the answer to her question. But surely there's no net change in requirement of ATP so its still as a whole is catabolic. I'll edit my answer.

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text:
I accidentally plated DH5a E. coli and left them in the 4℃ for 12 hours, if I put them into the 37℃ incubator will they start growing?
I started ligation of my constructs late last night and was super tired, since I used all of my ligation mixture I would rather not have to re-ligate.
I did my transformation procedure as follows:

Thaw on ice
Add 5 µL of ligation mixture and mix
Leave on ice for 30 min
Heat shock at 37℃ for 5 minutes
Add 500 uL of LB broth and mix
Incubate at 37℃ for 30 minutes
Spin down at 5000 rpm for 5 minutes and remove supernatant to re-suspend in 100 µL of LB broth
Plate on ampicillin agar plates with glass beads

Made a really dumb mistake and left them in the 4℃ instead of 37℃ overnight. Is this salvageable?

Go ahead and try it. Wat do you have to loose? My guues would be that it shouldn't natter.

If these were just regular E. coli that you were growing up, then it wouldn't matter too much - they would certainly grow a lot slower on the plate than you might otherwise expect compared to a plate put in the incubator straight after plating. The inhibitory cold and dry in the fridge means that they tend not to survive as well as they do after normal incubator growth and then storage.
However, as these are transformants and you only did a short recovery step of 0.5 h (~1 replication cycle), there's a good chance that you have lost the plasmid, so is quite likely that you won't get much growth because you have a selective ampicillin. I would be dubious about any that do grow as to whether they have the plasmid or not.
Much better to make a fresh ligation and try again after some rest.

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text:
How do you determine the structure of a molecular recognition feature (MoRF) after it binds to itself?
I have a protein that has a roughly 10 residue chain that registers at a MoRF in MoRFPred and MoRFchibi. I have the PDB files to correlate the known structure of the protein with the site of the MoRF; it checks out as the 10 residue chain sits in a known intrinsically disordered region (IDR) of the protein.
I need to find out what structure (⍺-helix, β-sheet, or other) that the MoRF exhibits when the two copies of the protein are bound together via the MoRF.
How would I go about this? Is there any way to do it computationally online?

I would suggest doing it using ColabFold, the colab implementation of alphafold2. The screenshot from the configuration shows you how to try to predict complex structures.

I'd give the whole sequence of the proteins and not just the fold, in the hopes that the results are more realistic.
Note that it is a computational prediction and not necessarily perfect. It may have difficulty with a disordered region, and it may also be that your region of interest is unstructured even in complex.


text:
What is (are) the species responsible for Lake Natron red waters?
Lake Natron is an alkaline lake located in northern Tanzania. It is known for its red waters, as seen on this image:

Natural-color Landsat 8 image showing Lake Natron in March 2017. NASA, public domain.
However, there seem to be some confusion around the species responsible for this color. Many sources state it is due to "cyanobacteria". Some sources have mentions of "spirulina" and "algae":

the lake actually derives its color from salt-loving microorganisms
that thrive in its alkaline waters. Spirulina, a blue-green algae with
red pigments, passes its pigments along to the Lesser Flamingoes that
feed on the algae and raise their young here. (NASA Earth
Observatory)

I know there is a lot of confusion around the word "spirulina", as the main species constituting the food, Arthrospira platensis, is not classified in the Spirulina genus. I also know there is a lot of confusion between "cyanobacteria" and "algae". Then comes another challenger, from a source already cited, but saying a different thing (emphasis mine):

Small, salty pools of water can fill with blooms of haloarchaea—salt-loving microorganisms that impart the pink and red colors to the shallow water.
(NASA Earth Observatory)

So I'd like to ask: what is (are) the species responsible for Lake Natron red waters?
Is it an algae? A cyanobacteria? A haloarchaea? And if so, which species? References to scientific articles containing biological analyses of the lake waters are very much welcomed.

Welcome Jean-Marie. Excellent first post here, impressively researched.

@JiminyCricket. Thanks! I have a few years of experience asking and answering on the Stack Exchange network, mostly at Earth Science SE. ;) Happy to join Biology!

As far as I can tell, no one has looked (at least not indexed on PubMed) in Lake Natron specifically for algal/bacterial/archaeal species. Lake Natron is a Soda Lake (high pH and high salinity) and is part of a chain of similar lakes in the East African Rift Valley, so there have been some general studies of these lakes, but mostly looking at zooplankton and flamingoes.
It seems that this is likely to be caused by members of the genus Dunaliella. These are strongly halotolerant green algae which have the ability to survive and thrive under salt conditions that most other species do not tolerate. This genus is cosmopolitan, so found world-wide and was first noted for turning evaporation pans for salt-making pink/red. The most likely species is Dunaliella salina, which is large and (apparently) turns more red than other species in the genus. The red colour is from accumulation of carotenoid pigments in the chloroplast and conversion of the green pigments into redder ones in response to extreme UV and saline levels.
There is another species that is possible called Salibacter rubra, but this has only been identified recently in Spain, so I don't know the global distribution.
I found a chapter The Microbiology of Red Brines1 by Oren in Advances in Applied Microbiology, 2020. to be of interest on this topic. As was Chapter 18 - African salt lakes: distribution, microbial biodiversity, and biotechnological potential2 in Lakes of Africa  which had this to say in the section on halophilic eukarotes (sec. 18.6.5)  (with my emphasis in bold):

Filamentous fungi of the genus Hortaea, Wallemia, and Aureobasidium, the yeast Debaryomyces, and microalgae of the genus Dunaliella and Asteromonas can be found in saline systems [65]. In addition, Dunaliella is the most ubiquitous microalgae in hypersaline environments [66]. It is a unicellular, eucaryotic, and green halotolerant microalga. The genus Dunaliella comprises almost 30 species, throughout fresh, marine, and hypersaline waters. The species Dunaliella salina is the most cultivated among the genus Dunaliella, for its high concentration (up to 10% of its dry cell weight) of β-carotene. When salt lakes dry out, salinity increases. These changes help the species D. salina to produce large amounts of β-carotene, which can turn the color of the lakes into red color.

You might want to check out references 66, 67 and 68 in that text, which are:

A. Ventosa, A. Oren, Y. Ma
Halophiles and Hypersaline Environments: Current Research and Future Trends, Springer (2011), p. 410

M.A. Borowitzka
Dunaliella: biology, production, and markets
A. Richmond, Q. Hu (Eds.), Handbook of Microalgal Culture: Applied Phycology and Biotechnology, John Wiley & Sons, Ltd., Chichester, West Sussex (2013), pp. 359-368

A. Hosseini Tafreshi, M. Shariati
Dunaliella biotechnology: methods and applications
J. Appl. Microbiol., 107 (1) (2009), pp. 14-35

Refs:

Oren A. The microbiology of red brines. Adv Appl Microbiol. 2020;113:57-110. doi: 10.1016/bs.aambs.2020.07.003. Epub 2020 Aug 17. PMID: 32948267.

Hafsa Yaiche Achour, Sid Ahmed Saadi,
Chapter 18 - African salt lakes: distribution, microbial biodiversity, and biotechnological potential,
Editor(s): Mostafa El-Sheekh, Hosam Easa Elsaied,
Lakes of Africa,
Elsevier,
2023,
Pages 501-525,
ISBN 9780323955270,
https://doi.org/10.1016/B978-0-323-95527-0.00009-9.

This looks great! I will check the references you provided asap. Thanks!

@Jean-MariePrival Thanks. Most of those refs are likely to be paywalled unless you have some academic access.

The chapter in ref 2 is available and contains some nice information: "High archaea densities in hypersaline environments can be observed with the naked eye due to the bright red, orange, or purple coloration. Archaeal strains are a promising natural source of carotenoids, which is why some water in hypersaline lakes such as the Magadi Lake in Kenya are colored bright red." Later: "These changes help the species D. salina to produce large amounts of β-carotene, which can turn the color of the lakes into red color."

And finally: "These water bodies are pink, orange, or dark red. This coloration is due to the presence of halophiles archaea, carotenoid-rich prokaryotes, and the microalga D. salina." So it looks like it could be both archaea and Dunaliella. I also kept looking on my side and found this article specific on Lake Natron https://doi.org/10.1038%2Fs41598-019-39935-3 which states: "Haloalkaliphilic archaea start to bloom under high salinities and give the lake an orange-pink-red appearance, especially observed in the upper (northern) half of the lake." With no reference to support this claim.

@Jean-MariePrival Refs 18-20 and 22 in your linked paper might be worth following up. Seems a lot of the literature on this is in books rather than papers, which makes it a bit more impenetrable.


text:
Will the sex ratio of chickens become favourable to females?
As male chickens are killed after birth in mass farming. Will being male become a negative evolutionary trait, that is to say will females become more likely as male death is more likely.

Also a follow up, does the chicken industry use selective breeding to increase the chance of females?

What do you mean by "the chance of females". I suppose you are still talking about sex-ratio. In all cases, please ask your follow-up questions on a separate post. Note that chicken are a GSD (Genetic Sex Determination) species. Just like other birds, they have a ZW sex chromosome system. There is therefore not much that can be done to bias the sex-ratio. Related post: Do males of all sexual species have Y chromosomes?

Any chick has a mother and a father. Hence, the contribution to the gene pool of the following generation comes at 50% from males and at 50% from females. As a result there is no selection for biasing the sex-ratio among its offsprings. The variance in reproductive success might be higher among males than among females but that does not matter for the sex-ratio.
This is known as Fisher's principle. In the terms of W.D. Hamilton (Extraordinary sex ratios), copy-pasted from wikipedia:

[..] given the condition that males and females cost equal amounts to
  produce:

Suppose male births are less common than female.
A newborn male then has better mating prospects than a newborn female, and therefore can expect to have more offspring.
Therefore parents genetically disposed to produce males tend to have more than average numbers of grandchildren born to them.
Therefore the genes for male-producing tendencies spread, and male births become more common.
As the 1:1 sex ratio is approached, the advantage associated with producing males dies away.
The same reasoning holds if females are substituted for males throughout. Therefore 1:1 is the equilibrium ratio.

To my knowledge, only kin/group selection can yield a sex-ratio different from 1:1 to be an ESS.

Let's assume all male chicks are killed, and all eggs are fertilised by some other population of males, not influenced by the farmed hens. Say for example a certain chicken (a female in this case) is more likely to produce females by some genetic mutation. Then chicks of said hen will also have a greater chance of producing females. This hen will have more female chicks than the average hen. Hence over time the sex-ratio tends towards females.


text:
Pronunciation of technical vocabulary
All knowledge of biology I have comes from online and after speaking for a brief period of time with someone from a biological education it is very clear that I have no knowledge of the pronunciation of all the necessary jargon I have learnt.
For example: Chordae Tendineae, Squamous Suture, Loop of Henle
How have people overcome this issue? I have searched online for pronunciation videos and whatnot, but very often they pronounce them differently to each other and it is impossible to know who is correct. I also do not want to learn the IPA for English.

"I also do not want to learn the IPA for English." - Yet this is clearly the best way to pronounce words where 1) you can find an IPA transcription for the word, and 2) you can't find a suitable audio source.

What's wrong with an online medical dictionary? E.g. hĕn′lē (sounds like HEN-lee), or  [kor´dah] (pl. chor´dae) (L.) Pl. sounds like CORD-eye. I learned by hearing other people, so no other good suggestions. Eventually patterns emerge, esp. because of all the Latin.

Don't know if it'd work for you, but I handle it by basically doing all my technical communicating in writing.

Come on guys! Let's use some 21st century technology.

I would try to use the audio features of dictionaries on the internet.
Ideally you would buy a good scientific dictionary app with audio pronunciation for your iPhone, just as I have for foreign language dictionaries on mine. However, the scientific offerings I found on the app store did not fit the bill (there may be something hiding there that I missed), so I looked on the web, and what I found was:
The Science Dictionary
Now this appears not to be an actual dictionary itself, but an aggregating site. So when I searched for Chordae I got several options, mainly from Wikipedia, which did not provide audio pronunciation, but also from
Dictionary.com
which did — for both the singular (which is what is the listing) and the plural.
Dictionary.com doesn’t always provide audio pronunciations. When I tried squamous suture there was only a definition — but there were individual pronunciations for squamous and suture.
(Loop of Henle was also there with pronunciation. Perhaps going directly to dictionary.com is your best bet.)
Rider and caveat
As @JM97 mentioned in a comment, some entries in the free dictionary have audio pronunciation, although others do not. There may be other on-line facilities that do likewise. However with any on-line dictionary one should ask oneself how reliable is the pronunciation; and it is difficult to tell because one doesn’t know how it is done in the way one does with a primary dictionary with a print reputation.
For example, take the pronunciation of the word "caeca". The open dictionary pronounces this in a classical Latin style with a hard C, whereas dictionary.com. pronounces it in a mediaeval Latin style with a soft C, the way it is pronounced by scientists I know who dissect it.

The free dictionary by farlex http://www.thefreedictionary.com/ is also very good.

@JM97 — I couldn't see any audio on the free dictionary.  Did I miss it somehow?

http://www.thefreedictionary.com/loop+of+henle see this example

Its a site which gives definitions from a large number of dictionaries including dorlands medical dictionary. just like science dictionary but the user does not have click an additional click.

@JM97 — Yes, I see there is an audio pronunciation for Loop of Henle, but not for Chordae Tendineae, which is why I had assumed it didn't support audio. (Also the app). I guess it varies. Another point is how reliable the pronunciations are. Compare the pronunciation of "caeca" on the open dictionary and dictionary.com. Better add a caveat to my answer.

It is also worth noting that in my experience, "accepted" pronunciation may vary quite a bit between biological disciplines and who a given specialist might have learned it from! For terms stolen directly from Latin, for example, there is often considerable variation between people about whether they should be pronounced in a Classical Latin style, an ecclesiastical/medieval Latin style, or (most commonly) an Anglicized style that sounds about right to an English-speaking biologist.
Names like the Loop of Henle are also tricky because they require you to know that they're anatomical structures named for particular people (in this case, Friedrich Gustav Jakob Henle), work out what language the person's last name is from, and then guess at what it is for an English speaker. English speakers do not always get this "right" on the first go, and weird pronunciations not infrequently result from confusions like this.
Personally, I wouldn't worry too much about it--I might just run pronunciations by a friend first and check to make sure I have them right there if I felt embarrassed.

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text:
Human brain is superior than till-date computers, yet why we perform poor than computer?
Scientists found storage capacity of synapses by measuring their size.
They found, on average, a synapse can hold about 4.7 bits of information.
This means that the entire human brain has a capacity of one petabyte.
This is same as about 20 million four-drawer filing cabinets filled with text.
Striving around this stuff, I have a weird doubt crawling in my mind and it goes like this: Why is it like this that a human whose brain has got a capacity to store 1 petabyte, i.e. 1 million GB, cannot fully remember even the contents and information of a simple textbook which on the maximum would not be more than 1 GB, when he has to appear in exam? Why is storage capacity not directly linked with remembering capacity?

@Ankur you haven't understood my question properly. I advise to you to kindly go through the question once more.

Maybe we need a RAM upgrade? :)

Three reasons:
Most of the storage capacity of the brain is used to store implicit knowledge. For example how to move your body, how to automatically recognise objects or how to hear and parse your language. These things may seem easy to you, but they require a lot of learning. Just look at how clumsy five year olds still are, they have been training how to use their body for five years straight and still skin their knees all the time.
Humans are actually able to retain huge amounts of information, both in their short term memory and in their long term memory. We know this because there are savants and hyperthymesia. It is likely that these abilities are actively suppressed in neurotypical people.
Because it is just not evolutionary adaptive to remember every detail. Intelligence depends on weeding out unnecessary information. If normal people learn huge amounts of information they often rely on memory abilities that are evolutionary adaptive, like the ability to remember a path from landmark to landmark.

To answer your question, we store information in a different way than a computer would, the storage happens in the connections in between neurons and it's not a perfect copy of what the environment presents, but rather certain features, I'll give you a classic example, read this sentence:
Tahkns for Cnotriubtng an asnewr to satck ecxahgne

You can understand the sentence even though the inner words are scrambled because it is not stored as a carbon copy, but rather as similar things attached to meaning within semantic and syntactic rules which in turn are stored as well, the recall and storage happens through a network in parallel, so the storage and the recall happens as you read it.
To give you a numerical example of how limited our brains are in terms of data as we know it, there are around 26,000,000,000 (26 billion) neurons on the Neocortex ( the number will probably go up-down in the future, but let's just use it for reference and assume it is only used for vision), your vision has 260,000,000 (26 million) cones and rods, where light is registered. If you were storing the information pixel by pixel in neurons alone, you would run out of space after just 100 frames. The synapses though  are a different matter,a conservative estimate is around 150 trillion: 150,000,000,000,000, and if you were storing perfect copies, you would run out of space after ~ 580,000 frames, or 4 movies.

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Cerebral/Cerebellar Cortex versus Deep Nuclei
I have troubles using the terms Cortex & Deep Nuclei, and 'Nuclei' in general. From what I understand, a brain has '3 matter types in accordance to anatomical locations'

Superficial Grey matter - This is called the 'Cortex (Cerebral or Cerebellar Cortex)'

Middle White Matter
This is called 'Cerebral/Cerebellar White Matter' separating the cortex and deep nuclei

Deep Grey Matter, deep to the cortex -This is called 'Deep Nuclei' (for e.g.: Basal Nuclei of the Cerebrum, and four deep Cerebellar Nuclei (dentate, globose, emboliform, and fastigial))

Quote from the book that helps me to jump into this conclusion or summary:

'The brain, like the spinal cord, is composed of gray and white
  matter. Gray matter - the seat of the neurosomas, dendrites, and
  synapses - forms a surface layer called the CORTEX over the cerebrum &
  cerebellum, and deeper masses called NUCLEI surrounded by white
  matter... (Saladin 2015, Unity of Form & Function, McGrawHill)

Thank you!

What exactly are you confused by?

Hi Jam. Thanks for your reply. I'm wondering that if we call anything brain's grey matter in any location 'nuclei' (Cerebral Cortex, Cerebellar Cortex, Basal Nuclei, and dentate, globose, emboliform, and fastigial are all nuclei)

In comparison, it's called 'cortex' in the superficial area, and 'deep nuclei' in the deep area. For example Cerebral Cortex & Cerebellar cortex are grey matter in the superficial part of the brain. For deep nuclei, examples are basal nuclei (in the deep part of the brain, deep to the cerebral cortex).

I'm mostly concerned of how we call grey matter in 3 different situations: (1) In general, either in superficial or deep brain locations (2) On the surface of the brain (3) Deep part of the brain, deep to the cortex & white matter. Thank you!

"Cortex" is a more general anatomical term for the outermost layer of a structure. It applies to both the cerebral and cerebellar cortex gray matter, as these are gray matter structures on the outsides of their respective parts of the brain.
Nucleus is also a more general term; in the context of neuroanatomy, it refers to a cluster of cell bodies. Typically we refer to CNS clusters of cell bodies as nuclei, and in the periphery they are called ganglia, though there are some odd exceptions like the basal ganglia (you referred to the basal nuclei in a comment, which are the same structure, but are almost always called the basal ganglia, whether or not that term is the 'best' term).
White matter, in general, is nervous tissue that appears white because it is primarily "fatty" from the lipid bilayer of cell membranes and consisting of processes like axons (especially myelinated ones) and dendrites rather than cell bodies.
I'm not aware of anyone who would call the cerebral cortex (or cerebellar cortex) a "nucleus": these areas are not shaped as clusters of cell bodies, they are shaped as sheets. The distinction is really only an anatomical one, so it makes little sense to use "nucleus" in a more general term to mean "nervous tissue consisting of cell bodies": we already have a term for that, "gray matter," and in most cases interesting to science we are talking about something more specific anyways.
I'd further say that these distinctions are really only gross anatomical ones, as one can have cell bodies outside white matter, and areas of gray matter like neocortex also include a mostly-white layer of axons and dendrites in "layer 1" but we still treat this as gray matter with respect to the whole brain.
Ultimately it doesn't matter: don't take anatomical terms more seriously than the things they are describing.


text:
Neuroscience - A neuron with two types of synapses (electrical and chemical) at the same time
I learn that the nerves from the Peripheral Nervous System can carry signals from and to other organs of the body.
I'm wondering if A Single Nerve carries
1) Only chemical signals
2) Only physical signals
3) Both of them at the same time
4) It can occur either of the case below, either (1), (2), or (3)
Thank you!

Homework questions (or homework like questions) are off-topic here unless you have shown your attempt at an answer (for more information see our homework policy). Can you show your attempt at an answer?

When you say "nerve" this refers to a bundle of axons from many different neurons, but I suspect what you actually mean is a single neuron.
Synapses occur at the ends of axons. Almost all axons branch to some degree, so there are many of these endings, and they fairly exclusively end in chemical synapses.
However, the actual propagation of the signal down the axon is electrical.
Electrical synapses ("gap junctions") also occur in vertebrate nervous systems, but they do not necessarily involve the ends of axons contacting dendrites of other cells. Rather, electrical synapses tend to be between dendrites of different cells, or between a dendrite and the somatic region of another cell, or between axons of two different cells.
The neurons that make electrical synapses also typically make chemical synapses in other places - sometimes they make these electrical synapses with the same cells they make chemical synapses with.
Note that these rules are different in invertebrates, and in general the distinction between axons and dendrites in invertebrates is more complicated.
There are other places besides the CNS with gap junctions/electrical synapses as well, including in the enteric nervous system and in the heart.

Draguhn, A., Traub, R. D., Schmitz, D., & Jefferys, J. G. R. (1998). Electrical coupling underlies high-frequency oscillations in the hippocampus in vitro. Nature, 394(6689), 189.
Galarreta, M., & Hestrin, S. (2001). Electrical synapses between GABA-releasing interneurons. Nature Reviews Neuroscience, 2(6), 425.
Traub, R. D., Bibbig, R., Piechotta, A., Draguhn, R., & Schmitz, D. (2001). Synaptic and nonsynaptic contributions to giant IPSPs and ectopic spikes induced by 4-aminopyridine in the hippocampus in vitro. Journal of Neurophysiology, 85(3), 1246-1256.


text:
When is it useful to give a vaccine post-infection?
I was reading this answer about the rabies vaccine, which explains that giving the vaccine post-exposure is effective because the vaccine stimulates a more rapid immune response than the infection itself would. From this, I inferred that the vaccine buys the immune system enough time to combat the virus, which would otherwise cause death before a natural immune response could fully develop. My question is: When this principle apply to other vaccines? My instinct is to say it would fail in the case of faster-acting diseases such as measles or the flu, because a natural immune response would have already been triggered before the vaccine could "teach" the immune system to fight off the pathogen.  Is that correct?  If so, would it mean that if we could figure out a way to get more vaccines to be effective more quickly (say, within hours or days, instead of the weeks or longer many vaccines take to induce even partial immunity), post-infection vaccination would be more generally effective, assuming the vaccine was administered as soon as possible after exposure, instead of rabies being a rare exception?
On the other hand, what about with dormant infections?  In the case of HPV in particular, I’ve read that vaccination is ineffective once the virus is already inside cells, as the immune system can’t target it there. That would seem to suggest that vaccines generally work by triggering an immune response that prevents a pathogen from entering the relevant cells in the first place.  And yet, the shingles vaccine prevents illness from a virus that's already infected the person and is dormant. So why does that work, but post-infection vaccination against HPV wouldn't?

The short answer is that it's always better to prevent an illness than to treat it. The only time this doesn't hold true is when prevention is riskier than the illness.
(I'll not address HPV, as it's a seperate question. Feel free to post it as such.)
For every vaccine, medication, or anything else we put into our bodies, a risk to benefit ratio exists. If the risk of putting something into our body is high, we don't take it unless the benefit is greater than the risk. E.g. We don't usually eat spoiled fruit, but if you are starving and the risk is death, the risk of eating spoiled food is low compared to the benefit (it might save you from starving.)
The risk of death from rabies is as close to 100% as it can be without being 100%*, so the risk very, very high, and the benefit is life. So the risk of post-exposure vaccine, which (especially with rabies immune globulin) has been found to prevent rabies once exposure has occurred, is very low comparatively.  The history of post-exposure rabies vaccine goes back to 1885 and Louis Pasteur. It was mostly because of the period it was developed in that the vaccine was given post-exposure. Most vaccines don't work post-exposure. Look at Ebola, a deadly virus. There's no post exposure vaccine, just treatment, which is not guaranteed, and much more expensive, and of limited availability, than vaccine.
We are far from those days. Modern vaccines are much safer than some earlier vaccines were, so, again, prevention is better a cure.
The significant medical problem following exposure to influenza virus aside from the usual (and very uncomfortable) illness is pneumonia. Even with treatment, roughly 30,000 people will die in the US alone  from complications of influenza every year (some years are worse than others). Depending on the year, many more will be prevented by pre-exposure vaccination (in the US, 6 million in 2022-23 according to CDC estimates.)
Usually, people cannot pinpoint the exact moment of exposure to a virus, and by the time the illness becomes known, it will be more difficult to treat than to have prevented it, so the "what if" scenario you propose would result in treatment failures and illness. A vaccine is relatively simple; it's not like we have sensors available to warn us of an exposure, and if we had tests available, to take the test every time we're exposed to people would be impossible, both financially and because of human behavior. Once the virus has started to cause symptoms, eradicating the virus is treatment, not prevention, and doesn't work uniformly. The risk to benefit ratio of this approach is untenable, for rabies, for influenza, for measles, even for chicken pox.
Shingles occurs when your natural ability to fight the virus is decreased. When this occurs, the dormant virus gets reactivated and starts to replicate and spread. This will result in shingles, sometimes, disseminated shingles or sometimes (rarely) something worse. When the vaccine works, it does so by boosting immunity to the virus so that it can be kept dormant and/or destroyed before it causes shingles.
*To my knowledge, there have been only ~15 known survivors of rabies (once symptoms started) so far in the world. Which should illustrate why treatment is less desirable than prevention.

Interestingly I came across a paper a while ago that looked at seroprevalence in humans (not sure if it was this one: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC8254041/) but it implied, as does the linked one, that rabies infection is much more common than suspected and largely survivable so long as it doesn't progress to the classic rabies neurological symptoms.

@bob1 - I answered another rabies question, and became aware at that time that there are people in remote areas where there is a high incidence of rabies that have IgG to rabies virus. At the time, it was unexplained, but clearly, there was exposure. So, yeah. But, I've now had PEP three times, and I wouldn't have it any other way!

Yes, I'm not in a rabies country, but I'm very glad the vaccine exists and works as PEP. It might have been your previous answer that made me aware of the incidence. Route of exposure could explain it, though there are those couple of intriguing cases where there's no evidence of bite. Though I guess there's still risk of contact with nervous tissue via inhalation.


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What is the name of this biting bug?
I just was bitten by a bug!!! It is 3mm long and has large pincers. Here is a photo:

I live in Colorado. Any help identifying it would be great.

It's a larva of a green lacewing (Family Chrysopidae). Yes, they can bite hard but you're not its intended victim and they're not only harmless but beneficial as they're aggressive predators of aphids and other soft bodied plant pests. I can't be specific to what species of lacewing as they look fairly similar.
Another larva that looks very similar to yours.

Source from BugGuide.net
What the adult looks like (remember,they're an entire insect family so there are differences but not to most people).

Source also from BugGuide.net

Wow! Thanks for the quick identification. Glad I caught it and released it ;)

That's rather interesting. The larva itself seems terribly threatening, while the lacewing looks cute!


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Has any research linked gestational period and exhibited mourning behavior? If so, how?
While reading the National Geographic Answer Book (2016ce), I was reminded elephants have a gestation period of a little less than two years.  Mourning behaviors for these creatures may include "sleep disturbance" and "increased stress." Chimpanzees and Japanese macaques may also show similar behaviors (gestations of 8 months and 171 days, respectively.)
The bottlenose dolphin has a gestation period of about 12 months.
Are you aware if the walrus and the white rhinoceros mourn the dead in their lives?  Both of these animals have long gestation periods.
Are any animals with long gestation periods not demonstrating these behaviors?
If any research has been conducted to link gestation or 'mothering' times to mourning behavior, how was this research conducted?  If not, how would you imagine we may best conduct this research?
Thank you.

How would you connect these? And what makes you certain to have "correlation and not causation"?

Welcome to SE Biology. Please finish reading the [Tour] to find how this site works and the Help on asking questions. You ask ask "us" how we would conduct research on a topic of interest to you, but we require posters to do their own research before asking questions. First, you might state that you have searched for "correlation of gestation period with mourning behaviour" and found nothing. However, more to the point, you need to show you have thought about the problem of determining when correlation implies causation. You will find this on the internet.

@Chris Hate to argue in the comments, but not all science is done in the lab. observational and correlation studies are legitimate science and can be quite useful in biology and other sciences. Especially when it comes to fields like wild-life biology, ecology, and animal behavior; where experimentalists are probably in the minority. Also, like, the vast majority of epidemiology research is based in observational studies, though it's maybe more common for epidemiologists to engage in both observational and experimental research, given the opportunity.

The field of study you're looking for is called comparative thanatology. Start with literature search on Google Scholar or another research repository, and use what you find to narrow the focus of this question.
I can think of some counterexamples already, but that doesn't mean there's not a trend. Domestic dogs seem to mourn each other, yet have relatively short gestation. Several Corvid species display what appear to be mourning behaviors, but how would compare compare egg incubation times with mammalian gestation?

Questions like this make me worry about the proper scaling of the measurements. After all, time could be viewed as relative to a) expected lifespan, b) expected fertility, c) body size, or more complex measures. "Year" or "month" are human created measurements, for all that they may have certain natural correlates, and do not represent non-human animal experiences of time. The proper unit of measure is not obvious without a supporting literature.

You don't state it, but presumably you mean higher animals, mammals, birds etc.. Creatures like ragworms have a phenomenally long gestation, but not much by way of a brain to process attachment and grief. What's your area of interest?


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What type of mold is this and is it dangerous to human and pets? How can I get rid of mold?

I put the in bathroom and this is what I got after 4 days.
Can you tell me what type of mold that is, is it dangerous to humans and animals and how did I get rid of it

This method isn't useful for the purpose you want because there are mold spores everywhere all the time and there's no getting rid of them.


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Differential Gene Expression
I have a matrix two gene expression datasets, one from 10 normal individuals and ten from diseased individuals, which are normalised log fold change values--, i.e., already processed.
My question is: how do I obtain a list of differentially expressed genes?
That is the up and down expressed genes. I was thinking of getting means for each group and then sorting the average fold change values would work, but I am not sure...
Thanks.

Note that there is also a bioinformatics stackexchange site: https://bioinformatics.stackexchange.com/

It hard to figure out what sort of data you have. qPCR? Let me give you an example with tools involved on how I get differential genes for RNAseq data, that is whole transcriptom and for mouse that transcriptom that's about 23000 genes.

Step 1. Map raw reads to the reference genome.
Step 2. Count number of reads within the feature. Feature can be several things, but lets just take protein coding feature (feature == gene).
Step 3. Do differential expressions analysis

From step 2 you get a matrix of counts.
      sampleA_rep1 sampleA_rep2 sampleB_rep1 sampleB_rep2
gene1      2             5        20             23
gene2      10            12       50             50
gene3      20            23       4              6
..
geneN

The packages that I use are limma and/or edgeR. You can find them on bioconductor Both packages take raw counts and give you topTable of DEG, default is top 100 genes.
I recommend reading edgeR or limma manuals, there come with great explanations as well as examples. Also A scaling normalization method for differential expression analysis of RNA-seq data is original paper that explains TMM normalisation (mention a bit more in depth below)
Two things; one you need to filter genes with low counts, convert to cpm() counts per million before filtering as this takes library size (depth of sequencing) into consideration. two calcNormFactors() will do TMM
The hardest thing (I understand) in RNAseq is to get total RNA output for each sample. Because a treatment will effect the number of transcripts expressed in that sample, increasing or decreasing amount of transcript, which give you unbalanced counts. You won't know just by looking at raw counts if geneA has less counts because it's been down regulated OR because that sample after treatment has more transcripts present in the cell and geneA simply has less coverage, due to other transcripts also getting reads now. I can't really explain that too technically because I don't understand this myself, but TMM normalisation takes care of this and  gives you an estimate of degree of variability.
I know that I'm not giving that much info on actual differential testing, which was your question. edgeR has exactTest() which is Exact Tests for Differences between Two Groups of Negative-Binomial Counts , which might help you, but you might have normalise your counts differently as mentioned above. The general idea (I understand) is you need to fit a model (lmFit()) and then do the test eBayes()
Other

Degust this is very nice tool that uses limma and edgeR in the backend, but simplifies a lot of things as well as gives you nice interface to work with your data. You upload raw counts! in the format shown above
DESeq one of the other rather popular R packages that can also do differential gene testing

There is software for this, but if you want a very primitive way to analyze it, do a t-test (which is basically looking at both the difference in means and the STDEV of each group)

At least suggest reliable software. 2) This primitive suggestion is actually a very bad idea, it ignores many of the crucial features of this type of data including compositional problems (which actually make this entire field of differential expression a bad idea generally), the overdispersion and count nature of the data.


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does small different between 16s ribosomal DNA leads to diffrence into protein products?
I have bacterium isolate after BLAST of its 16S  the different between its and the similar seq. on NCBI was 4 nucleotides my question is, does it lead to difference between whole genome seq. or I will obtain the same seq. consequently. does this also leads to  the same protein products from them.

The 16s ribosomal DNA does not code for any protein, it codes for the 16s ribosmal RNA, which is part of the (small subunit of the) ribosome.
The differences will most likely not affect the ribosome much, but to be sure you'd have to check the exact position of the mutations in a reference structure and see if they are at important positions (i.e. binding of the Shine-Dalgarno sequence)


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Identifying the Pheasants
I took a few photos at the Aviary of Hong Kong Park. The photos included some pheasants, and the below shows 4 of them. What species are they? I am not sure if 1, 2, 4 are actually the same species or not. Unfortunately on the official website http://www.lcsd.gov.hk/en/parks/hkp/birds_collection.php they are not included in the bird list, perhaps they are newly introduced.


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Is there a good protocol for bilateral decortication in rodents?
I was thinking about using bilateral decortication in rodents (preferentially mice) to study the roles of certain brainstem areas in cognitive tasks in the absence of cortex. However almost all studies I could find with bilaterally decorticate rodents date back to at least the 1980s if not older. Is there a specific reason for this? Also, do you guys know of any relatively recent protocol for bilateral decortication in rodents?

Probably seen as a bit ethically questionable these days, without a particularly keen need/reasoning. Also techniques are preferred to be able to temporarily turn parts of the brain on and off, for example with optogenetics, DREADDs, etc.

@BryanKrause - this is akin to answer in comment. Insert a few links here and there and (at least) I would be happy with this answer. Otherwise you might wish to delete the comment?


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How to choose which animals to study when designing evolutionary neuroscience research project
I am currently writing a reseach project for a scientific initiation and my idea would be a literature review identifying all empirical physiological and anatomical evidence for the presence and characteristics of a specific neural circuit in different animals, and possibly come up with insights about the development of said neural circuit during evolutionary history. My problem would be: which criteria should I use for choosing the animals to research? My focus would be to find the highest ammount of evidence possible for the presence or absence and development of this neural network across the evolutionary tree, so I was thinking about choosing animals that could represent certain taxons and also ones that have been extensively studied before; I wish to include both vertebrates and invertebrates such as mollusks to investigate the possibility of convergent evolution. My main problem is that I am a medicine student and although having some prior experience in neuroscience research, I have no formal experience in evolutionary biology. So I accept book recommendations as well as scientific papers on the topic that would help me. I also accept feedback on my idea, what do you guys think, is it too broad of a topic, too ambitious? I would have a year to complete the research and try to publish it as a paper, and the advisor I wish will help me is specialized in comparative animal neurobiology. But first I need to write a good project.
Thanks in advance.

What "specific neural circuit" do you have in mind?

I would suggest starting with a relatively simple neural system that is very well characterized; the obvious suggestion would be C. elegans. It's not clear to me why you would need invert/vert both, you will probably see significant relatively interpretable variation (including convergence) even in related worms. One might consult e.g. https://www.ncbi.nlm.nih.gov/pmc/articles/PMC6748829/. How workable all of this is depends heavily on your answer to Bryan's question.


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Why does the posterior region and the right ventricle of the heart are not well represented in standard 12 leads?
When assessing an ECG we divide our 12-lead into four main contiguous lead groups: inferior, lateral, septal, and anterior. Regions not well represented on a standard 12-lead are the posterior region and the right ventricle, So if there is an infarction in these regions how we can determine it ?

The answer should have source or reference and not the question and this is known in ECG that lead l , aVL,V5 & V6 looking for lateral region of heart ...ect as shown in photo above.


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How the glycogen accumulate intracellularly in the reversible cell injury?
As we know in reversible cell injury as the most common cause is ischemia decreases oxidative phosphorylation which decrease ATP & increase anaerobic glycolysis which depleting glycogen !!? But one of the intracellular accumulation in addition to lipid , hyaline, pigments and calcium is glycogen So how this occurs ?


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What is the mechanism behind sympathic effusion of serous cavities due to nearby abscess?
Patient with left perinephric abcess which does not ruptured have reduced breath sound that is mostly due to plural effusion revealed by CT scan which was sterile. I found this called sympathetic effusion , Why it named as this and what is the mechanism behind it ?

The left perinephric space is adjacent to the left subpleural space so inflammation can spread from the former to the latter.
Diaphragmatic inflammation then results in a left exudative sterile pleural effusion. It's called sympathetic since it's not infective.
https://radiopaedia.org/articles/perirenal-space
https://emedicine.medscape.com/article/1979032-clinical#b4


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Why placental progesterone suppressed or drop before onset of labour?
As we know during pregnancy predominant hormone is progesterone which is at early pregnancy about first 3-4 months produce by corpus luteum  of pregnancy together with estrogen after that placenta can make its own progesterone with estrogen till birth but for initiation of labour estrogen go up and progesterone should go down because it prevents uterine contraction. Does anyone know mechanism of suppressing of progesterone and rising of estrogen before onset of labour ?
In other word mechanism of initiation of labour ?

Pregnenolone -the precursor of progesterone is principally converted to estradiol instead of progesterone.Thus the oestrogen:progesterone ratio increases prior to parturition.

Fetal cortisol stimulates activity of placental 17 α hydroxylase/17, 20 lyase (CYP 17) enzyme, which catalyzes the conversion of pregnenolone to estradiol.

Confinement of the foetus in the uterus stimulates the stress responses.

Stress leads to a rise in foetalACTH. The rise in ACTH causes production of cortisol from the foetal adrenal glands. The rise in foetal cortisol leads to an increased oestrogen:progesterone ratio by upregulating aromatase enzymes, which convert androgens to oestrogens. Cortisol also reduces progesterone production.


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Would asymmetrically positioned eyes impair vision?
Assuming one eye is 5 cm further down compared to the other, would that somehow change perception if the brain has long enough to deal with that incoming information? For example, if you were born that way, would you be able to do depth perception and binocular vision just as well?

Welcome iwab. You'll notice the "do your research" stipulation in the [ask] section of the [help]. Please show that research. For example, if an eye were positioned in the maxillary sinus instead of the orbital socket, would there be an optic-nerve to connect it to the brain? What about muscles? What do your researches into stereoscopic vision lead you to conclude in this case?

If the eyes have the same distance then they geometric effect is the same. if the eyes were on top of each other, they would have less depth perception information when the person turns and leans their head, and they would have a greated change in depth perception when they are running. It's probably tested on robots, not animals.

Would you be able to do depth perception and binocular vision just as well? This is hypothetical question. Hypothetically, the answer is yes.
The human brain has an amazing ability to adjust to changes to an individual's physical body.  As evidence of this, consider the PD (pupillary distance) when you were born to what it is now.  You have the same brain (in all likelihood), but that distance has undoubtedly increased significantly.  Your vision continues to work exactly as designed.
This amazing flexibility is not limited to homo sapiens.  Some species of actinopterygii are born with eyes on both sides of their head, but develop into fish with both eyes on the same side of their head.  Their ability to hunt does not diminish after this change takes place.


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Sequence identity and coverage in multiple sequence alignment of proteins
This might be naive since I am very new to the field, but I wonder about the difference between sequence identity and coverage in multiple sequence alignment of proteins. I imagine the calculation would be simple, but I couldn't find strict definitions of these two terms and how they are calculated. Can somebody possibly provide an example showing how these two terms are calculated (preferably sequences with gaps and the terms not being 100%)? Many thanks.

I've never heard the term "coverage" used as in this area, and sequence identity is obvious until you start to deal with sequences of different length, when it is not. So the answer is that different packages will calculate on the basis of what seems reasonable to them. If you are asking about a particular MSA program you need to consult the documentation — or look at the results and work out for yourself what is going on. In extremis you could post an example here. Of more practical interest is "similarity" — scoring for similar amino acids. See posts on this list.

In high-throughput sequencing applications, we will often refer to "coverage" as the number of unique sequence reads (queries) that align to the same region in a reference sequence. This alignment problem is a subproblem of the alignment domain referred to as "local" alignment, and coverage makes sense in this context as a measure of how many local alignments of short sequences map onto some location in a much larger sequence.
However, the multiple sequence alignment context that you are referring to is likely a bit different, where you do not have a reference sequence and instead try to find some maximally good alignment of a set of query inputs. This is sometimes called the "global" alignment problem. Here you could plausibly compute the number of other sequences that have aligned residues at the same positions as some focal sequence. That would be a bit like "coverage".
But I agree with commenter David that measurements like similarity are probably more useful in practice- they will certainly be highly correlated with such a "coverage" measure in any reasonable example I can imagine.
Minor update: pairwise identity is a very simple measure that just measures the number of residues in one sequence that are aligned to identical residues in a second sequence. I'm surprised that you can't find anything on this, but here is one highly technical discussion that probably covers a bunch of cases that you don't care about.


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Antigen transfer from baby to mother
When considering immunity in babies, the main mechanism that supports their undeveloped immune system is the transfer of antibodies from mother to baby via breastfeeding.
Case #1: both baby and mother got sick with a virus; the mother then produces large amounts of antibodies that are passed to her baby via breastfeeding.
But what if the mother (for some reason) does not get sick while the baby does?
Case #2: just the baby got sick.
What are the possible scenarios for supporting the baby's immune system in that case? It looks like the number of relevant antibodies is low in the mother.
Another mechanism could be envisioned, namely, the baby transfers antigens to the mother when breastfeeding, which stimulates the mother to produce antibodies and pass them back to the baby. Is this mechanism real? Is it efficient in any sense as compared to Case #1?
(scientific references would be appreciated)

Are you suggesting that in case#2 that the mother has not been exposed to the babies' pathogen? (i.e. is the baby isolated from the mother, more-or-less precluding breastfeeding) Or are you suggesting that the baby has been injected with some sort of antigen for which there is no infectious disease present?

@ARogueAnt. - I believe the OP is asking, what if baby was exposed to the pathogen but mother was not? (Say, baby went to visit grandmother, gets virus from visiting cousin. Baby's infection would precede mother's, if mother did catch baby's virus.) It's a reasonable hypothesis, no "isolation" or "injection" necessary.

To the OP: You say, "When considering immunity in babies, the main mechanism that supports their undeveloped immune system..." Can you support that the child's immune system is "undeveloped", since your question hinges on this? Please provide support in an edit to your question.

@anongoodnurse is it important whether the baby's immune system is developed or not for this question? I'm only asking about the possibility for the mother to do the dirty job of making antibodies while not being sick herself.

If that's what you're asking, then edit your question too include that precise request.


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is pepsin able to break down proteins into amino acids in human stomach?
I'm trying to find out if pepsin can break down proteins into amino acids after they are converted to peptides. Is pepsin able to separate amino acid monomers from the peptides?
can you please show me the actual reaction happening in the process?
I'm a 10th-grade student so please describe it as suitable for my knowledge.
thanks in advance.

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.


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Sodium-potassium pump (Na+/K+) time limit, endless without action potential?
As written in Wikipedia:

The sodium–potassium pump mechanism moves 3 sodium ions out and moves
2 potassium ions in, thus, in total, removing one positive charge
carrier from the intracellular space.

I know that thus the cell (neuron) is excitable, and due to an action potential, the ion concentrations will change since their flux is opposite as what the pump does.
But what happens if there is no action potential, and the pump tries to work forever until no sodium ions are in the cell anymore?
Presumably, this won't happen in reality but in theory then the pump could not keep the cell polarized?
Or are there other effects like leak currents that contribute, and why exactly that issue won't happen?

This won't happen, so if you want to ask about a hypothetical you're going to have to be very very precise.

Even if there is no leak or other currents there will be an equilibrium condition in which the ATP hydrolysis by the pump could drive no more ions due to the electric potential. Actually there are condition in which the pump can work in reverse, synthesizing ATP in the process.


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Troubleshoot bacteria fixation for SEM
I fixate bacteria on titanium coins for 1 h i 2.5 % Gluaraldehyde. Then wash 3 times in ddH2O. Then 10 min in each solution of Ethanol 10 %, 30 %, 50 %, 70 % 90%, 100%. Finally soaking in HMDS in 15 min, then take out the HMDS and let dry overnight. Sputtercoat with Ag/Pd. I know I have bacteria on the titanium surface from the beginning but I dont see anything in SEM. What am I doing wrong?


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Pseudomonas aeruginosa biofilms grows stronger after 95 % ethanol treatment
I used 95 % ethanol to disinfect Pseudomonas aeruginosa biofilms from titanium discs and compared it to disinfection with water and hydrogen peroxide. The strange thing was that when examining in SEM, roughly ten times more bacteria was present in the ethanol treated samples than the water treated samples hydrogen peroxide treated samples showed no bacteria. Metabolic activity assay showed however more bacterial activity from the water treated samples than the ethanol treated, hydrogen peroxide showed least. Anyone have any theory why ethanol seamed to increase the bacterial biofilm compared to water? Ethanol are regarded as an effective disinfectant on bacteria.

95% Ethanol does not work as a disinfectant. You need to dilute it to around 70% to be effective.

@Chris while that is true, wouldn't you still expect it to be better than water?

@Chris is right. ~70% ethanol is the way to go.
From a discussion here :

The effectivity of ethanol as e.g. desinfectant or antiseptic agent depends on the concentration of ethanol-water-mixture: An ethanol percentage of 50-80% destroys the cell wall/membrane of bacteria by denaturing their proteins and dissolving their lipids (effective against most bacteria, fungi and some viruses; ineffective against bacterial spores). Therefore, the ethanol has to pass the bacterial membrane/wall to get into the bacteria - if you use 100% ethanol instead, the bacteria get 'sealed' and they will survive... An other mechanism is the high osmotic pressure of ethanol/water-mixtures; and the 70% has the highest one.

By using a high ethanol concentration, you're effectively creating a layer of coagulated protein on the surface of the biofilm, "sealing" the bacteria in below that top layer.
As to why water seemed more effective than alcohol, I would venture a guess that some of the bacteria from the biofilm was flushed off. Maybe check the water for bacterial count to verify? :)


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Which cancer cell lines do not increase glucose uptake in comparison with their wild type analogues?
I heard that some cell lines which already have a high glucose influx do not increase it upon altering to a cancer phenotype. However, I cannot find any supporting references.

Where did you hear?

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.


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Does the kidney regulate sodium balance or total body sodium
Imagine the following situation. You have a person who initially eats 10 mEq/day of salt. He then, at t=0 begins to eat 150 mEq/day of Na, and will continue to do so, because you're forcing them to, forever. My question is about how the kidney responds to such a situation. There's two options.

Option #1: The Kidney ensures sodium balance (sodium intake = sodium excretion). In this situation, the kidney regulatory mechanisms ramp up Na excretion until it equals 150 mEq/day. But since this takes a couple of days, there is a bit of time in which intake > excretion and so total body Na rises. Because of osmoregulation, more water is drunk to balance the increased total body Na, and so the person's weight rises.
Option #2: The Kidney brings total body sodium back to normal. In this situation, the kidney regulatory mechanisms ramp up Na excretion, but it briefly ramps up excretion to more than 150 mEq/day (excretion > intake) to excrete out the increase in total body sodium that built-up. In this case, the weight gain is temporary.

See a graph describing the two different responses.

The situation is inspired by Chapter 40 of Boron & Boulapep's Physiology textbook. The reason for the question is that Boron states that Option #1 is correct, whereas my teacher in class said differently  ¯_(ツ)_/¯ .

What specifically does your book and instructor say happens? I suspect they differ a bit from the scenarios you've crafted in ways that make them quite the same, instead of in conflict...

I phrased the question to my teacher exactly as I have done so here. I asked, assume you have a person and you have them increase their salt intake consistently - a hamburger every day is the example I used - what would happen to their weight: would it go up then down, or up and stay constant at the new high value.

And what does the book say?


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How Difficult is Artificial ATP Synthesis?
One way of making ATP requires:

A membrane (would probably have to be a phospholipid bilayer)
A difference in H+ chemical potential across the membrane
ATP synthase anchored to the low-potential side of the membrane
Some reasonable concentration of ADP on the low-potential side of the membrane

Now, I know ATP has a short half-life, but if we wanted to run some step of a biological process that requires ATP, would it be possible to create an artificial ATP synthesis "cell" that would convert ADP to ATP for use by enzymes in solution?
If not, then are those four steps sufficient? If so, then which step is the hard part? Thank you.

Welcome to SE Biology. As a new user we expect you to do us the courtesy of at least finish reading the [Tour]. You have not done so. Please do, and look at the Help on asking questions. Questions that relate to hypothetical situations can generally only be answered by opinions. Creating cells is usually considered in the domain of God, or perhaps evolution. What precisely do you mean by “is it possible to create an artificial ATP synthesis cell”? What exactly is the biological problem you are faced with? Questions here are expected to fulfill the latter criterion.

Hello David. When I said "cell" I wasn't referring to a biological cell but an electrochemical cell. I do not know why you would have interpreted as the former. Regarding what I meant by "is it possible to create an artificial ATP synthesis cell?” I meant precisely what I wrote. My problem is synthesizing large amounts of ATP in vitro. My question does conform the the tour guideline which you can see because it's already been given a non-opinion answer.

I’m not much the wiser. What’s an electrochemical cell? Sounds like something in the chemical domain. And why do you want to generate large amounts of ATP? The fact that someone posted an answer, means nothing, especially as the answer is in terms of a biological cell with a lipid bilayer membrane,and amounts to no more than touching on the main features of the biological system. If I wanted to generate ATP I’d use a single reaction that performed substrate-level phosphorylation, but I don’t answer questions which are unclear to me. Btw we use @johndoe type references for alerts to replies.

If your just trying to mass produce ATP there are several known production systems, why do none of those work for you?

The four steps you listed are the basic requirements for ATP synthesis, and a simplified artificial ATP synthesis system can be created using these components. However, there are some additional factors that need to be considered in order to create a functional system, such as the stability of the lipid bilayer, the efficiency of ATP synthase activity, and the availability of energy sources to generate the proton gradient.
One of the key challenges in creating an artificial ATP synthesis system is the efficient reconstitution of functional ATP synthase in the lipid bilayer. ATP synthase is a large, multi-subunit protein complex that is composed of both membrane-bound and soluble components, and it can be difficult to reconstitute the complex in an active form in vitro.
Another challenge is the generation of a stable proton gradient across the membrane, which is required for ATP synthesis. This can be achieved using different energy sources, such as light, electrochemical gradients, or chemical reactions.

there are many TP production methods that do not need ATP synthase and use enzyme cascades instead.


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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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What kind of bird did I hear at three AM in Germany
I woke up at three AM from some sort of industrial noise outside. In addition to the noise I heard some creaking like from a stuck door. After the initial noise had ended, something that sounded like birds calling continued for at least 30 minutes.
The most common call sounded like staccato tweeting that lasted for about a second and increased in tone height. This call I heard many times. There were also some slower, non staccato beeps but no continuous singing like you'd hear from a blackbird.
I'm in southern Germany, in a city near a park. The area is brightly lit. It could conceivably be bats, but in my experience they don't tweet loudly and they dont make a staccato noise.

Try to listen on line a nightingale, maybe it was a far away one and you could hear only the most characteristic notes which are similar to what you describe. Are you south or north of Munich (roughly south of the city there are not breeding nightingale).

It could be a bird that is normally not out and about at that time, but also disturbed by the industrial noise.

A bit North of Munich. I think birds decide whether to sing by whether it's light out, so I think it should be a bird or bat that normally sings early, even if the noise woke it up.

Bats are quite out of question. You would not be able to hear most of their sound, and they definetly wouldn't remotely remind you of a blackbird.
Re: nightingale, try http://www.xeno-canto.org/species/Luscinia-megarhynchos - but I have my doubts.


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What is the best method to check for differential gene expression in large dataset? I want to identify aging differentially expressed genes
My data is RNAseq data and I want to know which all genes (which I am looking for) are differentially expressed with age. I have tried the Kruskal-Wallis test and fold change methods. Do you have any better suggestions?

Someone here may be able to answer your question, but if not I would try SE Bioinformatics, which might have been a better bet in the first place.

A standard pipeline for DGE would be Salmon for (pseudo)mapping + DESeq2 for statistical analysis.
Salmon is one of a set of modern, fast and accurate mapping software. Requires a transcriptome to be defined.
DESeq2 is a mature R (Bioconductor) package which can handle reasonably complex designs, including time series, but not mixed models. The vignettes are pretty thorough. It can handle data from pseudomapping software as well as data generated using more traditional methods, e.g. Star + FeatureCounts.
Using a pre-made RNAseq statistical package is a much better idea than trying to do, e.g. a Poisson/Neg.bin GLM manually for each gene (even though ultimately this is what DESeq2 does).
(agree this question is a better fit for SE Bioinf)


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Is this an insect? Which one?
I found this animal, which I suspect it's an insect, on the floor of a kitchen in Spain some days ago. It's about 8-10 mm long. I don't think I've ever seen it before. What is it?

This looks like a silverfish (https://en.m.wikipedia.org/wiki/Silverfish )

It is a silverfish. It is a 'basic lineage' in the insect class. You'll find it here on OneZoom.org.
Silverfish are considered household pests although I don't think they spread disease nor cause much damage (thanks to @WilliamBligh for his comment). They typically live in the canalization (bathroom, kitchen) or anywhere that is wet enough. In a building, they may be hard to get rid of as they can easily move from one apartment to another. You have to keep your place very clean and eventually use toxins such as Permethrin or Deltamethrin.

While they are considered a "household pest", they are to the best of my knowledge not harmful. Even contamination of food is, again to the best of my knowledge, undocumented (but sometimes assumed). If they are not present in large numbers, I would advise not to poison them.
Edit to add: pyrethroids are hardly "specific" toxines.

You're right. Thanks for your comment. I edited the answer accordingly.


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My Queen ant has laid eggs but has not dropped her wings, is this ok?
My Queen ant has laid eggs but has not dropped her wings! I caught this Queen in England, I've had her for three days and she's laid eggs already but I was expecting her to to drop her wings first as she needs the wing muscles to feed her brood. I don't yet know the species of the queen but I'm suspecting it's a camponotus queen but I'm not sure, the queen is 1cm long and 4mm wide, the gastor is bigger than the thorax and she is black. (I haven't got a recent photo of the queen because I am currently away on holiday and will be for the next month. (My mum is still at home though so that's how I know there are eggs and the queen still has her wings)).Is there a species of ant that never drop there wings or don't drop them for a long time?

You might have better luck asking some of your questions on a forum specifically for ant-keeping. Not so much that your questions are off-topic here, but we have a relatively small community and it might be quite some time before someone with the relevant expertise comes across these questions.


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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.


text:
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.

Tarsal cuffs maybe? The insect equivalent to "feet" is tarsi and tarsal claws- that is, the final leg segment that makes contact with the ground and the claws that grip it. Sorry this is so late.
https://www.amentsoc.org/insects/glossary/terms/tarsus/#:~:text=The%20tarsus%20is%20the%20final,Telotarsus


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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.


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

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

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

Alpaida versicolor
Alpaida veniliae
Alpaida carminea
Alpaida bicornuta
Alpaida rubellula

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

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


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

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

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

Milliseconds to days.


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Are animal whiskers a useful monitoring tool?
I have come across studies that have used stable isotope values from seal/sea lion whiskers to infer foraging ecology. However, I am also aware that whiskers can fall off and then grow back. Is there a way to determine the age of a specific whisker, to verify the timeline stable isotope samples represent? E.g. if a sample is taken every x mm that correlates to the foraging ecology y years ago.

theforestecologist might have a great answer for this. For me, one wait is to feed harmless radioisotope carbons during a specific observation time. If you can regress one way to another the rate of whisker growth, then I would be probably feasible

Whiskers have very similar science to other hair follicles. You need to specifiy the types of whiskers, rat, seal or tiger whiskers?

I expect that vibrissae growth rate would depend on the species, the type of vibrissae, time of year, etc. One paper that comes to mind is Zhao & Schell 2004, which estimated growth patterns of harbor seals (Phoca vitulina) using labeled tracers:
https://www.int-res.com/abstracts/meps/v281/p267-273/. I am sure there are other papers that have studied pinniped vibrissae growth rates. I recommend a more specific search using Google Scholar.