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What global habitat classification systems are there?
I am an IT guy, not a biologist or ecologist, but have ended up with (roughly) the task "given this Latitude/Longitude pair, return the 'habitat' at that location". The points of interest in practice are likely to come from Europe or North America, but could come from anywhere. All data points will be terrestrial, not marine. What habitat means isn't fully defined, and I'm looking for a best-practice answer.
At this point I thus have a question: "what are the well-regarded global habitat classification systems?". A basic search leads me to the IUCN Red List Habitat classification 3.1. From there, I find some nice recent research work by Jung et al that attempts to map the world onto an IUCN habitat type by combining data from a Koppen-Geiger climate classification system, from Copernicus land-cover data and from an ecoregion+biome classification coming from the WWF. This classification has had a number of iterations, but currently seems to have 867 ecoregions in 14 biomes. The resulting map is at approx. 100m resolution and maps on to 47 of the IUCN habitats.
To my amateur, all of this seems great, but is it the state-of-the-art? Or are there other habitat classification schemes out there? Where should I be reading?
Many thanks indeed for any help and apologies in advance if I have broken any rules of the site - this is my first post.


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What are these structures in blood from a nosebleed?
I decided to look at some blood from a nosebleed (not mine) under the microscope and took some pictures of the results. There many large orange structures interspersed within the blood cells and I'm wondering if someone knowledgeable can tell me what they are? Are they skin cells? What are the fibres inside them?

The first two images are x400 and second one is x100.

Could you estimate the diameter?

In the first picture it is roughly 100 microns if I use the RBC as a unit of measurement.

This is a mere guess: You see accumulations of airborne fibers (dust) that got caught by secreted lipid droplets within your nose.

source: Stack Exchange

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

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How do schools of fish get enough oxygen?
Fish get oxygen from the water running over their gills, during which up to ~80% of oxygen is removed from the water. In schools of fish, how do fish swimming behind other fish get enough oxygen in this case? It seems counterintuitive that the volume of unenriched water is repleted with oxygen within the fraction of a second before the fish following behind attempts to gain oxygen from it.
Thanks.

Welcome. Where did you read the claim 80% of O2 is harvested? It seems like an overestimate. Further, fish move through the water and water is in motion as well - how would that affect O2 saturation?

This beautiful paper from 1958 cites a dissertation by van Dam, 1938, which I couldn't fully access, for the figure of 50-80%, and I see "80%" referenced widely in study resources online.  It is presumably part of a basic textbook.

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


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Is there a reliable reproduction rate for adenoviruses?
I was trying to find a R0 for adenoviruses but the best I could find was a study done in a bootcamp in China that infected 375 people.
This states that the R0 was found to be 5.1:

The basic reproductive number was 5.1 (95% CI: 4.6 to 5.6)

Is there any other more reliable method that covers more data?

I expect it depends on the virus strain, social conditions etc. As such, this isn't a very good question.

@Fizz Well obviously, that is why the one example from the study isn't a very relevant answer - the R0 can vary, but all viruses generally have a mean R0 or range, that is what I am trying to find out.


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Help with histological cell identification in colon
I have scans of a histology slide of tissue of colorectal cancer and its surrounding tissue. This slide has been stained with the typical Hematoxylin & Eosin (HE) staining then unstained and stained again with two immunehistochemical markers, one for each B- and T-Lymphocytes. Those markers are CD3 for T-cells (which appears red) and CD20 for B-cells (which appears brown). The two scans were then aligned so that it is easy to identify a lymphocyte on the immunehistochemical stain (IHC stain) and find that exact cell in the HE stain.
After looking at these scans for some time I noticed, that there was a cell type that, to me, looks indistinguishable from a lymphocyte on the HE stain but clearly expresses neither CD3 nor CD20 when referencing the IHC stain. These cells are mostly found inside colon epithelium or cancer. Below are two images (one in the HE stain and one in the IHC stain) with two example cells marked with red arrows.
My question is what type of cell this is? Could it be some sort of precursor cell or sub-class of T- or B-cells, that expresses neither CD3 nor CD20? Is it even an immune cell at all?

There are immune cells, such as monocytes, macrophages, and dendritic cells, that don't express CD3 or CD20. I'm not enough of a histology expert to identify them on this slide, though.

Monocytes are, by definition, only present within blood vessels, macrophages have a large diameter with slightly uneven borders and a large amount of cytoplasm and dendritc cells aren't found inside epithelium to my knowledge. So I'd be very surprised if it was any of these cells.


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Electrochemical and concentration gradient confusion
I am learning membrane transport and learnt that there are 2 gradients that determine passive and active transport.

If the concentration of a cation is higher in the cytoplasm side (hence ions should flow out of the cell passively), but there is higher electrochemical potential in the extracellular side (hence ions should flow into the cell), which side should that ion flow if there is only passive transport?
I know there is Nernst equation for electrochemical potential, but I would like to ask whether there are equations for calculating potential caused by the concentration gradient? Since my thought is by comparing which has the higher potential difference, and deduce which gradient is dominant, eventually deduce the direction of the net passive flow of that ion.
Thank you

All you need is the Nernst equation and the voltage across the membrane.
The answer you get from Nernst is the equilibrium potential for the ion of interest. This is the potential at which the concentration and electrical gradients are exactly equal for that ion. If the voltage across a membrane is at the Nerst potential there is no net passive flow of the ion.
If the voltage is different then the Nernst potential, then the ion moves in the direction that would tend to move the voltage towards the Nernst potential. If the Nernst potential is -90 mV, the membrane potential is -60 mV, and you have a positive charged ion, the ion will tend to move out of the cell because this is what would move it closer to -90 mV.
You can't separate the chemical and electrical gradients and sum them up or anything, the specific way to combine them is given by the Nernst equation. The Nernst equation is not for calculating the electrical potential separately of concentration gradient, it's for calculating the combined result of electrical and chemical gradients. The way to know which way an ion flows is the way I've described it, not by determining which is "dominant".


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What is the correct nomenclature for expressing a genotype where a recombination event may occur?
Given an example punnett square:
 -------------------------
 | x   | A-b     | Y     |
 -------------------------
 | a-B | A-b/a-B | a-B/Y |
 -------------------------
 | A-b | A-b/A-b | A-b/Y |
 -------------------------
 | a-b | A-b/a-b | a-b/Y |
 -------------------------

What is the correct nomenclature to express the genotype prior to putting it into a punnett square? The parent's genotype looks like a-B/A-b x A-b/Y except I want to show that a recombination event can occur, yielding the third row in the punnett square table.
This is relation to my previous question about how to calculate recombination frequencies.

I would suggest "genotype".
A Punnett square is a visualization technique to help us think, it doesn't change the terminology.
If what you mean is the 1N genotypes represented in the rows/columns that combine to make the 2N genotypes, you might consider "haplotype" (haploid genotype). The haploid cells with those genotypes might be called gametes, but that refers to the cell and not to the genotype.
If you are specifically interested in those genotypes/haplotypes in which a recombination event has occurred (or not), you could call them recombinants or non-recombinants. In some cases you can encounter specialized terminology, for example in yeast tetrad analysis you encounter terms such as "tetratype", "nonparental ditype", "parental ditype". But those are not generally used outside the yeast field, I believe.


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How many coronavirus particles are in the body at the peak?
How many coronavirus (or other common virus) particles are in the body at the peak? How far it multiplies? Are there billions, trillions or other number of them?

This measure changes dramatically between viruses and infected individual but there are some estimates that have been done.

Bar-On, Flamholz, Phillips & Milo, Apr 2020, eLife
From MERS and SARS (similiar to the current coronavirus) we know that from each infected cell emerge around 10^3 virions and it takes them 10 hours from the infection of the cell. We cannot deduce that every 10 hours the magnitude of virions in the body increases by 3 because:

Not all virions infect a new cell
Some virions infect the same cell and the burst size depends greatly on the cell
Not all virions are viable or able to infect

Another paper from Sender et al. (Nov, 2020) that is still in preprint (this is a long time for Coronavirus related articles) estimates the number of virions in peak infection at 10e9-10e11.
This estimate was done by analyzing data from Rhesus macaques (a monkey).
Concentration of virions per tissue weight was multiplied by the tissue weight to get total amount of virions.
On the relevance of Coronavirus information
Since our knowledge of Coronavirus advances and changes all the time it is important to note the date and source of information. Also, to advance science many papers published preprint articles regarding Coronavirus. This articles are published before peer-review and should be taken with a grain of salt as every peer-reviewd paper gets many revisions before published to the public.


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Which enzymes use ATP?
It is well known that there are many enzymes which use ATP in their function, and any enzyme that work against an energy gradient need to have that energy supplied from somewhere, but just as well there are enzymes that simply work spontaneously and do not require such an influx of energy. However I can't seem to find any sort of list, however brief, that contains examples of enzymes that use ATP and ones that do not, or any indication of their prevalence. I could go one by one through each of the commonly encountered enzymes in the cell, but that would be tedious.
Is there any source where I could find what I am looking for? Or at the very least which of the major enzymes in the protein transcription/synthesis chain require ATP?

This would be such a massive list I can't see why you would maintain such a thing. It's possible someone will have an answer to how to extract it from a bioinformatics database, though.

I agree with @BryanKrause but both he and I gave examples of ATP-utilizing enzymes in our answers to a different question. This might be a starting point. However please finish reading the [Tour] and the help on asking questions here. We are keen on people showing evidence of having tried to answer their question themselves.

I just searched the Expasy Enzyme site for ATP using this search page. Will take you a while to look through them although there were fewer answers than I expected.

Thank you, David's answer was just what I was looking for. However I should have expected the daunting task of picking out a set of examples from that list. Any chance of being able to sort by a criteria like papers cited or times cross referenced, to a get some sense of importance?


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Can anyone ID this fossilized bone
Is this a fossilized backbone? If so then from what?
This "bone" is approximately four inches (10 cm) wide by two inches (5 cm) deep and two inches (5 cm) tall. The texture is rock-like. If it resembles any bone familiar to anyone let me know or take a guess and I'll look into it. I found it in North central Texas in the ground by a massive fruitless mulberry tree on the top of a hill. It weighs about what a 2x2x4 inch chunk of average run of the mill stone would weigh.

Welcome to Bio.SE. As written, this post -- in my opinion -- is unanswerable and will be closed. Please review the guidelines for asking [tag:species-identification] questions. You need to update your post with size(s), location, circumstances, etc. Given the unclear nature of your specimen, photographs from multiple angles with better lighting will be necessary. Describing the weight and texture might also be relevant. Please add as much detail as you can. Thanks

Please add a more specific location than "North America" Also note that "by a tree" is not helpful — please add environmental and geological information to the best of your ability. ——— In addition, in the future please proofread and edit your posts and make sure all the pictures are visible (you had inserted links into each other breaking most of them). Thanks!

Thanks for the updates. Please be very specific about the location you found it. North America is too coarse of a location -- ideally, we should know the exact location (or as close to it as possible). You should at least include the city. Knowing the ecology, natural history, and human history of the site you found it will also be relevant. If you know any of this please share. If not, then a specific location (and photo) would be best. Also, please provide more info about how you found it: What type of tree? on what type of property? How far down in the ground? Soil type? Etc...

It looks like just a rock, not fossilized bone.

You can put this on the earth stack to be answered. I suggest adding pictures of all sides of the object not just multiple pictures of the same side from different angles, specifically and broken areas are helpful. locality will also be helpful, right now it just looks like a rock.


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Do the primers used for Covid-19 test work with all new strains?
We keep reading news of new Sars-CoV-2 strains, some of them allegedly able to evade vaccine-induced Igs. That's not unexpected, specially for a single-stranded RNA virus. However, it seems to me that the tests (RT-PCR) are being done exactly as they've been since early 2020, all around the world.
Therefore, my question is about the primers used: how can we be sure that there are not new strains with sequences not complementary to the primers? Does the current covid19 test target highly conserved RNA regions? If that is correct, how do we know that those regions are highly conserved?

Just for completeness, these Q/A provided no relevant clue: What specific markers does a Covid-19 PCR test look for? and Are specific primers or detectors, or both, used in COVID-19 tests?.

Here's a good resource for anyone that wants to try answering: http://covid-19.dnageography.com/. This is the SARS-CoV-2 Mutation Browser, which, as of writing this, includes data on >82k mutations at >6k positions across >10k complete SARS-CoV-2 genomes. Conveniently, the data can be subset to look at position-specific mutation frequency across genes and regions, including the regions primed by the most popular diagnostic primers, like the primers from the CDC's diagnostic panel.

accompanying preprint: COVID-19 Variants Database: A repository for Human SARS-CoV-2 Polymorphism Data

I TRY to answer. First of all, no we cannot be 100% sure that the primers are effective against all strains.
However, the strains that are under media attention right now are due to variants in the spike protein, while RT-PCR primers are designed on different genes.
One WHO document (I do not know if it's the official guideline) targets RdRp (RNA-dependent RNA polymerase) and a gene called E gene.
While the document itself nor the paper on which it is based explicitly addressed conservation of the target genes, I found evidence for relatively high conservation of at least RDRP in this paper.
Personal opinion here: in looking for RT-PCR primers, you always need a balance between targeting genes that do not vary in the species, and targeting genes that may aspecifically amplify in different species (e.g. SARS-CoV).
Final answers:

how can we be sure that there are not new strains with sequences not complementary to the primers?

We cannot. However, as far as the variants media are talking about, we know that RT-PCR has some ability to detect them, since they have been sequenced as a consequence of some patient testing positive to RT-PCR.
We cannot exclude the existence of unknown strain. They could be identified by e.g. sequencing patients with COVID-2 symptoms testing negative to RT-PCR (and possibly testing positive to the antigenic test). I don't know if such cases have been observed.

Does the current covid19 test target highly conserved RNA regions?

Yes, the current RT-PCR based tests target conserved genes (although I wouldn't call the highly conserved, because of the caveat I expressed above as a personal opinion)

If that is correct, how do we know that those regions are highly conserved?

Mostly by sequence similarity. Regions that have the same sequence in all the individuals of a species and in related (or distant) species are conserved.

No, some strains contain mutations causing partial or complete failure of amplification of at least one of the test targets.
Here's an FDA bulletin on the issue from January:
https://www.fda.gov/medical-devices/letters-health-care-providers/genetic-variants-sars-cov-2-may-lead-false-negative-results-molecular-tests-detection-sars-cov-2
As the bulletin notes, most tests have multiple targets, so the test result remains positive with a virus variant even if one of the targets doesn't amplify.  In fact, the UK B.1.1.7 variant prevalence has been estimated using RT-PCR by looking for a one-target failure result.
Different tests use different primer sequences and even target different Sars-CoV-2 genes. That's a lot of different mutations that would have to happen to evade all detection by even one test, let alone the many different ones. If Covid-19 symptoms started showing up in many people testing as "negative", it would be quickly be investigated in more detail.


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Basis of enzyme nomenclature — pyruvate dehydrogenase
In the formation of AcetylCoA from pyruvate, why is the enzyme called “pyruvate dehydrogenase (complex)” when it involves the decarboxylation of pyruvate or the replacement of a carbonyl group by coenzyme A?

The pyruvate dehydrogenase complex catalyses the oxidative decarboxylation of pyruvate. A description of this is easy to find on the web (e.g. Wikipedia), but I would recommend the section in Berg et al. Biochemistry (online), as it is an authoritative book with a high reputation.

Pyruvate + CoA + NAD+ → acetylCoA + CO2 + NADH

In the overall reaction, shown above, pyruvate is being oxidized (hydrogen removed) by NAD+, which is reduced to NADH. Hence there is nothing surprising in the enzyme having acquired the description, dehydrogenase.
Before asking questions about enzyme nomenclature, one should consider the historical and scientific context in which names arose. This was general scientific ignorance of the overall metabolic pathways in which they occurred or even the direction in which the reaction normally proceeded. There is also the question of focus — here on oxidation of carbohydrates. The working names that arise — trivial names, as they are called — tend to stick: they are in the literature and everyone in the field knows what you are talking about. Eventually, when the dust has settled, scientific commissions assign systematic nomenclature. In this case there are three enzymes involved, so that each has its own systematic name:

Pyruvate dehydrogenase (acetyl-transferring) — EC 1.2.4.1
Dihydrolipoyl transacetylase — EC 2.3.1.12
Dihydrolipoamide dehydrogenase — EC 1.8.1.4

It can be seen that even the systematic name for enzyme E1 (which catalyses the decarboxylation) does not refer to the decarboxylation, but to the transfer of the acetyl group (to thiamine pyrophosphate and then to lipoamide), which is of more interest mechanistically.
Details of the reactions and alternative names corresponding to the EC numbers  (Enzyme Commission numbers) can be found either on the Queen Mary University site or Expasy.


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How are thoughts biologicaly generated?
I want to know how thoughts are biologicaly generated. I know that electrical impulses can trigger formed memories in the brain but what I want to know is how the electrical impulse is generated in the first place. I don't have a background in biology or neurology but I think it's probably one of the three options:

Impulses are generated randomly at parts where there are stronger connections.
We have loops in our neural network, electrical impulses stay and go around the brain to trigger memories and thoughts.
We are deterministic and only react to impulses from the outside.

Any reading suggestions about this topic are also highly appreciated. I want to understand the human brain a little more to compare it to artifical neural networks.

Hi! It's an interesting question, though I would caution you to narrow down your question, it's very broad. I also warn you to not confuse thoughts or memories with neur(ologic)al activity, this will run you into the hard problem of consciousness. Please also show some effort in trying to answer your own question; for instance, what exactly is it that you don't understand? You will have to show some independent effort for your question not to be closed or marked as a homework question.

An addendum: since neural circuitry is extremely varied, you will find many examples of all sorts of circuits, including those you list. For instance, there exist neurons with a pace-making activity that generate their own activity from within, as well as neurons that only respond to outside stimuli, such as many kinds of sensory neurons. And everything in between... including neurological dysfunctions. You MUST narrow your question and also be very cautious about comparing all of that to artificial neural networks (the analogy is full of common pitfalls and frequent misunderstandings).

Thank you for your answer! Although not posted as an answer your response did help me out. Do you have a resource suggestion where I can deepen my knowledge about different neural activities and differences? I know that artifical neural networks in the current state are completly different from how our brains work. I do believe though that a lot can be learned from our human brains for AI. I think my question can be closed now. And sorry again for the form of my question.

Please take a look here and try searching "how are biological and artificial neural networks different" in a search engine. There are many duplicate answers on psychology.SE and biology.SE and so many more answers and posts about it elsewhere. Good luck!


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Were nicotinic acid/amide or flavin nucleotides ever part of primary RNA sequence?
NADH and FADH2 redox reactions are built deep into our biochemistry.  For example, pyridine nucleotides are involved in >500 enzymatic reactions.  When we look at the structure of deamido-NAD+, it is strikingly symmetrical:

The same is true for FAD, a two-headed monster of a nucleotide

in which we can recognize the two heads, belonging to xanthosine (a purine nucleotide) and alpha-ribazole 5'-phosphate, the latter of which is produced from flavin in the course of B12 synthesis:

The phosphoribose can be swapped between alpha-ribazole 5'-phosphate and nicotinic acid.
Given such enticing hints, it is tempting to speculate that one or more of these nucleotides might be so fundamental because it was actually part of some ancient RNA primary sequence, giving RNA containing this code the power to do serious catalytic chemistry with acid and amide groups, powerful options for redox and methyltransferase activity and more.
I know this is a hard question to answer, but is there any evidence that weighs in, or could weigh in, on whether this happened?  For example, if these functional groups are too unstable in regard to intramolecular reactions on a strand of RNA, or if there is no way to envision a potential base pair that would be reliably distinguished from the known base pairings.  Or if RNA editing sometimes creates them in some organism.  Or if aptamers made from these subunits show improved catalytic possibilities.  (That doesn't seem so easy to look up, since many papers look for aptamers binding these as prosthetic groups, rather than incorporating them)  It would be useful simply to have a notion of what evidence for such an idea would look like.

Have you searched the literature relating to thoughts on the adenine nucleotide components of NAD and FAD? I am not familiar with this myself, but it clearly must be extensive. My own thoughts on the subject (probably off-topic here) do not include your suggestion of these having been "bases". Nicotinamide and flavin do not look remotely like the bases, have no obvious "pair", and are essentially later oxidising agents to the likely more ancient Fe–S electron redox centres. If they arose to supply the redox function which ribozymes lack, how might they bind to the latter?

Your thoughts shouldn't be off topic - if you can make a good case for late evolution of NADH and FADH2, then it would rule out the idea.  But to my eye, nicotinamide and 5,6-dimethylbenzimidazole don't seem any more different from purines and pyrimidines than those two are from each other.  (I'm not claiming to know what they would base pair with)

There are NAD capped RNAs [1,2] but that's not what you are asking, right?

There are two reasons that I think this question off-topic. The first is that although you have asked for chemical arguments, answers are likely to be matters of opinion. The second is that it does not really conform to the question and answer format of this site, but is more to the discouraged type "I have this idea, am I right?". The way I see of resolving this (because I find the topic interesting) is given in the following comment.

@WYSIWYG: They are useful for your answer, since they show that the NAD isn't excessively reactive in the context of a longer RNA sequence.

A way of resolving the problem would be to rephrase the question to allow a variety of answers, not just the one you have in mind. This would be in of the type "NAD and FADH have adenine nucleotide components which are not involved in the redox reactions — show structures — Why?" You could then answer your own question with your idea, which could be subject to the usual criticism in the form of votes and comments. I could add mine. This would be a bit hard on @WYSIWYG who has just answered your question, but he might consider going along with it.

@David: I think a question of that type would be called "unfocused", and in any case, is a different question.  How do I ask about whether the RNA world had this range of catalytic activity - without asking it?  I think this is already open to a variety of answers and can be answered in the negative as well as the positive.

As you will. I will refrain from commenting on your idea for fear of causing offence, but I do suggest that rather than focussing on chemistry in isolation you consider the evolutionary sequence of events that your idea would entail and attempt to find out what others have written on this topic.

@David: It sounds like you have some useful thoughts to contribute regarding the evolution.  I have trouble deciding what to make of such data.  Does the metabolic relationship with tryptophan mean that nicotinic acid evolved from it, or the reverse?  If methyltransferases act on nicotinamide and RNA, but they're not closely related, what does that mean?  I'm not sure how to get to a conclusion from this sort of information.

Mike. I'd like to discuss this sort of question, but  I have been rather busy over the last few months and am only just re-engaging with the list again in a small way. I am still quite busy and have some other questions here that I need to tidy up loose ends on. I would prefer to wait until I have time to do some reading in this area and then post a question that I feel can generate some good answers. (The great thing about being a scientist is that you never retire. It's just that there comes a time when people aren't paying you any more.)

There are naturally occurring RNAs that contain a NAD "cap" at 5' end (Bird et al., 2018; Wang et al., 2019).
I could not find any papers on natural RNAs with nicotinamide or flavin as a regular base. However, a study with synthetic oligonucleotides reports that NAD containing oligonucleotides can form stable triplex structures (Göckel & Richert,2015).
There are two studies from 1970s that explore base pairing potential of flavin. Raszka and Kaplan (1974) report that FMN can hydrogen-bond with AMP but not other nucleotides. They further explore this finding to show that FAD can form intramolecular hydrogen bonds. Scarbrough et al (1976) show that flavin containing nucleotides can form stacking interaction in addition to base pairing kind of hydrogen bonds. (Compare D-X bonds)

More recently, Julius et al. (2020) have shown that NAD and FAD can act as co-factors of primase to initiate DNA replication.
Overall, it seems plausible for FAD and NAD to be a part of nucleotide chains (more plausibly for RNA as they are already known to contain a variety of unusual bases). However, I cannot comment if they really did play such a role during cellular evolution.


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Have we reached the technology to 'drive' evolution artifically?
Previously our evolution was due to circumstance, we can trace back all our body features to overcome some 'challenge' our ancestors were faced with. Now, in the modern time, with that large progression of science and technology, do we have methods to drive an organism to evolve to suit some artificially induced circumstance?
Further, have new species been formed this way?

Do you consider dogs and wolves to be different species?

I hadn't known that came from the same lineage. It was quite an interesting thing to read about @jamesqf

Look up the term artificial selection.

Every species we've domesticated was the result of humans 'driving' the developments of desirable characteristics.  Humans have been doing that for at least 10,000 years.

I doubt that domestication was from the beginning a directed effort. Most likely it began as coecistence of species. There's also evidence that there was continuing gene flux between the "domestic species", such as dogs and horses, and their wild counterparts long after the domestication began.

For some species it absolutely was directed.  Still is.  The Bible has a story about an attempt to generate a desired phenotype in the next generation.  The methodology is rubbish, but it shows that people were thinking about how to purposefully control the characteristics of the next generation

Oldest parts of the Bible were written about the first millenium BC, i.e., about 3,000 years ago. If you have scientific references to actual "species" (rather then breeds) that evolved over this time, it is worth including it in the answer. On the other hand, the Bible mentions lots of the species that had been already domestic at the time of its writing.

Three different concepts humans have already invented and attempted fit your qualifications: Artificial Selection (domestication), Eugenics, and Genetic Engineering. Obviously some more distasteful in their application than others, but yes, we have created new species in these ways, in the sense that they are not able to reproduce with their "natural" counterparts.

Short answer, Yes.
As @swbarnes2 point it out, traditional artificial selection has been done in pets and agriculture for centuries. Nowadays this principle can be used to accelerate evolution on bacteria implementing in-vitro evolutionary algorithms

Let me first note that the term evolution applies to species rather than individual organisms (the OP says do we have methods to drive an organism to evolve to suit some artificially induced circumstance?) Evolutionary changes happen over many generations, whereas the changes in a single organism during its lifetime are not really evolution - its genome remains pretty much the same.
Secondly, the obvious answer given by others - the artificial selection requires some qualification: while dogs, horses, cows and other domestic animals have evolved into separate species due to their cohabitation with humans, it is not always obvious that such evolution was a result of deliberate human efforts. Both in terms of its deliberateness, but also in terms of whether any such continuous effort lasted long enough for a new species to develop. We can however meaningfully speak of deliberately developing new breeds of plants and domestic animals.
Finally, there are multiple (sometimes ongoing) experiments of evolution in various closed ecosystems, experiments at deliberate development of microbial resistance to antibiotics (aka morbidostat) and so on. (I will add the references later.)


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How to assemble three 60mer nts by pcr?
Good morning,
I am new to molecular biology. The question might be silly but i would like to know the answer. I have three 60mer single strand synthetic oligonucleotide. Namely Tag 1 - 3. My goal is to assemble these three together by pcr. So i designed two primers having overlap of 18nts from each tag 1 and tag 2 and then Tag 2 and tag 3 ( i mentioned the orientation of my primers). What is the best way to assemble. I read about PCR- based two-step DNA synthesis. But i would like to know more about this.
If you have any questions please feel free to post. Thanks and looking forward for your valuable answers
[]

In order to assemble your TAG oligos by PCR you will need to redesign your primers so they are complementary to the TAG oligos as DNA polymerases work by adding nucleotides to the 3' end of a DNA strand.
Your design would look like this :

Now with this design you won't be able to assemble everything together as TAG-3 complementarity to primer 2 only presents 5' ends to be extended, which your DNA polymerase can't do. So you can either order the entire reverse complement of your TAG-3 oligo or design a primer 3 that would anneal at the 3' end of TAG-3 to amplify TAG-3.
With all these parts, you should be able to assemble everything in a single reaction, but depending on your downstream process you may have to purify the product of interest as the single reaction will generate intermediates (1, 2, 3, 1+2, 2+3).

Really thanks for your explanation. So i have to redesign the primers that should share  complementary with tag-1 and tag-2 (not reverse complement right). I will assemble first tag-1 and tag-2 and then i will go for tag-3 with my product.

You should order the reverse complement of your primer 1 and 2. Let's say that your TAG-1 sequence is 5'-attgcctg-3' and TAG-2 sequence is 5'-TAATTGCC-3'. So far what you have ordered as primer 1 is 5'-ctgTAA-3' but you need the reverse com 5'-TTAcag-3'

Awesome. Once again thanks.

@Rengaraj If this answer addressed your problem, please consider accepting it by clicking on the check mark/tick to the left of the answer, turning it green. This marks the question as resolved to your satisfaction, and awards reputation both to you and the person who answered. If you have >= 15 reputation points, you may also upvote the answer if you wish. There is no obligation to do either.

I would suggest using Gibson assembly. I am not an expert but this is the kind of problem that it is designed to solve. Probably there is some way to do this by PCR but I expect that it will be quite finicky.
See this quote from that page (my emphasis):

Tip: “Stitching” Fragments Together using Oligos
When you need intervening sequence between two PCR products, one method is to “stitch” together several oligos. This technique is especially useful for introducing promoters, terminators, and other short sequences into the assembly and is used when the part to be inserted is too long to include on overlapping PCR primers (>60 bp) but too short to make its own part (<150 bp).
Please note that the way to design the “stitching” primers and the amounts of them to include in the Gibson reaction are different than with normal PCR primers. The details are published in ( Nat Methods 2010; 7:901-3 ).
Tip: Number of Fragments Assembled Simultaneously
Multiple fragments can be assembled in one reaction. However, some labs have observed a sharp decrease in success rate when assembling more than 5 fragments at a time.

Update
It looks like there are people who argue both that Gibson is impossible with short fragments or that it works fine. There is more information on this here and here and here. Due to the relatively complex nature of your reaction, I think that Gibson might save you time. But other people linked there do claim that PCR works better.
Update 2
GaelC's answer above has better/more specific methodological advice on PCR assembly.

I agree with Gibson assembly but i read that it is not suitable for short fragments such as 60-mers.

@Rengaraj see update

Exactly, these are the posts i went through. First i will check by PCR assembly, if it did not work i will order the Gibson assembly kit. And i have a doubt? the way i designed the primers are fine or correct? or should i have to change anything...

@Rengaraj I think that GaelC's answer is good if you choose to go with PCR.

NEB's tech support is fantastic. If anyone has the answer about Gibson assembly of shorter sequences, they would.


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Interaction of charged triphosphates with the ATP binding region in proteins
DNA-histone interactions involve positively charged amino acid side chains groups neutralizing the negatively charged phosphate of the sugar-phosphate backbone. Is the same true for ATP binding sites on proteins?
In most models I’ve seen, it looks like the base, adenine, is involved in the binding. But the triphosphate seems like it would bind specifically to positively charged side-chains. I Googled a bit, but I couldn’t really find any models depicting this. Does anyone know whether this is so?

Thank you for accepting my answer. I enjoyed answering your question — perhaps in more detail than you required — but the answers to questions on this site are meant to be generally useful, rather than a dialogue with the poster. I had a little spare time in which I improved the illustrations and changed their order. I suspect that the answer was not quite as simple as you expected, which is not unnatural if you are new to the field of proteins. The answer hints at the complex intellectual beauty of this area, which will well repay the further work required to explore it.

a bit low detail actually

Do tell me what you would specifically like more detail of and I'll see if I can help. Some indication of your scientific background might help me pitch things at the right level.

Summary
Many proteins that bind the nucleotide triphosphates ATP and GTP have an evolutionary conserved motif — the P-loop — that binds the phosphate portion of the molecule. This has the sequence, GXXXXGKS/T. Although the conserved basic lysine (K) is interacts with two of the positively charged phosphate moieties, other interactions involve Mg2+ ions (with which NTPs are always complexed) and polar, rather than charged, amino acid residues and the polypeptide backbone.
Detailed Illustration
Although the question asks about ATP, I have chosen the GTP-binding protein H-ras p21 as an illustration because the paper by Pai et al. describing it gives full details of the interactions of the phosphates, which are essentially the same for ATP and GTP. (Many other papers tend to show only ribbon diagrams of the protein.) Note also that the crystal structure was, of necessity, of a complex with the GTP analogue, GppNp. (GppNp is not rapidly hydrolysed, unlike GTP itself.)
In this case the evolutionary conserved P-loop that binds the phosphate has the sequence:
Gly Ala Gly Gly Val Gly Lys Ser

 10  11  12  13  14  15  16  17

A three-dimensional representation of the phosphate-binding region is shown below. The left-hand frame shows the P-loop and residues Thr35 and Asp57 without the nucleotide. The right-hand frame is the same but with GppNp and a magnesium ion included, and only the conserved residues of the P-loop labelled.

[Own work based on Fig.6 of the paper by Pai et al. using Jmol software and Protein Data Bank file 5p21.pdb.]
A two-dimensional schematic of the interactions is shown below. The side-chain interactions (and those from the magnesium ion) are shown as red dotted lines, whereas those from the backbone are shown in black. The conserved residues of the P-loop are indicated by blue text.

[Simplified redrawing of part of Fig.5 of the paper by Pai et al., excluding water interactions and interatomic distances.]
Reflections
Although positively-charged basic amino acid side-chains are involved in neutralizing the negative charges of nucleic acids and nucleotide phosphates, they are not the sole means of doing so, and in the case of the NTPs they are not the main means. Magnesium ions and polar residues (including the protein backbone δ+ve NH groups) can play an important role. To my mind this reflects two things. First, that life is dynamic; so weaker interactions are often favoured in enzymes because they can be easily broken as well as made. (NDP must be released from the enzyme after hydrolysis.) Second, and more speculative, the P-loop is thought to be very ancient as it is found in a wide variety of conserved NTP-binding proteins. A specific backbone conformation has been identified that facilitates this interaction. It could have arisen before all 20(ish) proteins specified by the genetic code had arisen, when the protein backbone could have played a greater role in interacting with small molecules. (This has also been suggested for backbone NH groups binding δ+ve sulphur in iron–sulphur clusters.)


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Why do huskies have markings above their eyes
[1.         Dogs in Antarctica pulling sleds

It's not just Huskies,  Many breeds - Rottweilers, Dobermans, Bernese - have them.  Others like German Shepherds may or may not have them as part of their coat color variation.

You may be able to get better answers at [pets.se].


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What is meant by 4 –12% or 8% SDS-PAGE?
I am reading a journal paper and I am looking at the materials and methods section. Regarding the Western blot method in the paper, I have come across the following statement:

Proteins were separated by 4 –12% or 8% SDS-PAGE and electroblotted to
nitrocellulose transfer membrane.

I am not sure what is meant by '4-12% or 8% SDS-PAGE' in the above statement. Do the percentage values refer to the percentage of acrylamide in the gel? Any insights are appreciated.

Do the percentage values refer to the percentage of acrylamide in the gel?

Yes. The 8% gel is 8 g acrylamide per 100 mL. The “4-12%” gel is a gradient gel, which are useful for separating proteins over a large range of sizes. Read more about the uses and formulations of gradient gels here.


text:
Evidences supporting that coronavirus (SARS-CoV-2) came from bats
I was trying to find evidence that the coronavirus (SARS-CoV-2) came from bats. So far I know that coronavirus shared 96.2% overall genome sequence identity with a bat coronavirus RaTG13 from horseshoe bats who identified as natural hosts for SARS-related coronaviruses which were the direct progenitors for the origin of SARS-CoV and also comparing the receptor-binding domain (RBD) of 2019-nCoV to SARS-CoV, the structure of the RBD allowed for the use of ACE2 (angiotensin-converting enzyme 2) as an entry receptor. I wanted to know that apart from these two pieces of evidence are there any major evidence that also suggests that indeed SARS-CoV-2 came from bats.

I think the current consensus is that SARS-CoV-2 didn't originate in bats, but rather passed through them from another species before jumping to humans. I don't believe that other species has been identified yet, but I may be wrong - this is from news reports.

@MattDMo Isn't the hypothesis the reverse? I mean the virus existed in bats, then was passed to another species, then to humans?


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Why is the start of my coding sequence ATG and not TAC?
I am engineering a set of genetic sequences and have come across a surprisingly basic point of confusion that seems to have fallen through the cracks regarding coding sequences.
The standard start codon is AUG, and since this is an RNA sequence, I would thus expect that the start of the coding sequence would then be TAC, since DNA is complementary to the mRNA transcript taken from it. Wikipedia and various introductory texts confirm this expectation, e.g., in wikipedia's article on the central dogma:

The ribosome reads the mRNA triplet codons, usually beginning with an AUG

Here's another example: in the Khan academy introduction to the genetic code, the codon illustration clearly shows an AUG in the mRNA sequence.
The coding sequences that I am composing into my design, however, are all listed as starting with ATG, rather than TAC, which would be the complement of AUG. I am certain the coding sequences are not wrong, since some are very well established parts like BBa_E0040.
Something seems to have gotten confused or elided in many standard explanations, then. Why do coding sequences start with ATG and not TAC?

This image from the Kahn Academy article 'Overview of transcription' might help:

Essentially, the sense/coding strand of the DNA encodes the sequence that is transcribed. The RNA polymerase binds to the antisense/template strand, for which the code is indeed TAC, but when it then transcribes this strand it is again complemented, giving us the AUG that is recognised by initiator tRNAMet (also known as tRNAfmet), as shown in this diagram from the Kahn academy article on 'tRNA and Ribosomes'.

(Note that this diagram is actually showing Met in the middle of a sequence, rather than the start. fMET binding looks slightly different.

Thank you, that is very clear. I'm surprised that this is not more clearly presented in more places, since everything I'd found either said nothing about which strand or else used terms like "template" and "non-template" that weren't clear about how they related to promoter location strands.

Yes! I think this is something that is intuitive to people who've been through the standard Biology training so it's often not explicitly explained to folks approaching SynBio from different angles. It's good to have a clear question here so that people can find it in the future.

When your start codon is 5'-AUG-3', it's clear that the template DNA strand should be 5'-CAT-3'
If you look at Noah's illustration, it's also clear that the RNA sequence is basically the same as the sequence of the non-template strand, and runs in the same direction.
People generally refer to the mRNA not in terms of its RNA nucleotides, but in terms of the cDNA which it could be converted to.  But they'll reported it in the direction that makes sense for talking about RNA.

I'm afraid that your answer contains the sort of mirror-language ambiguities that caused my confusion in the first place.  Which cDNA to convert it to, the template or the non-template? From @NoahSprent's answer, I now know that the template is the opposite from the strand with the promoter, but if I didn't have that answer to read, I would still be confused.

cDNA is double stranded molecule, so it contains the template and non-template strands.

Yes, of course: that's why there's the ambiguity that I was finding confusing.


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What is the thickness of the membrane if only alpha helixes are embedded of a transmembrane protien?

Given is the representation of a transmembrane protein. Calculate the thickness of the membrane if only alpha helixes are embedded in it. One turn = 5.4Å
Please read:
The reason I didn't submit my attempt is that, I thought because there isn't an exact and precise depiction of the protien molecule and also the question is vague so I hesitated. I counted the number of turns ie 7 but still there was a lingering doubt "what if I am wrong, I would make a fool of myself."
What I know:

Protiens basically bunch of amino acids bound by polypeptide bond.
Primary structure is the sequence of the amino acids.
The secondary structure us due to the Hydrogen Bonding between the Oxygen atoms having partial -ve charge and Hydrogen atom attached to the Nitrogen, within the backbone only. eg: alpha-helixes, beta-sheets, Turns, Loops
Tertiary Structure is due to the interaction between the different R groups in the amino acids. Like the disulphide bonds, hydrophobic interactions, H-bonds, Vanderwaal forces.
Quaternary Structure is due to the overall 3D structure which is result of the aggregation of the polypeptide subunits.

Please go easy on me, its my first time posting here and I am still a 1st year in college and 2nd Sem is gonna start next week.

Please finish the [Tour] to find out how this site works. Your question is incomprehensible without reference to external sites, which is quite unacceptable. It would also appear to be a homework question which, should you edit it so it is comprehensible by itself, requires you to show your attempt to solve it.

Welcome to the site. What research have you done on your own to answer this question? The Biology.SE community has agreed that questions that show little or no prior research effort are off-topic on this site 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. Unresearched questions may be subject to down-voting and closure. Please see our homework policy for more information.

@David Sorry, I should have included it in here too. It's my first time posting here.

The question begins by telling you that only the alpha helices are embedded in the membrane. This means that if you can calculate how long the alpha helices are, you can work out the thickness of the membrane it is embedded in.
Alpha helices look like spirals (similar to fusilli pasta shapes). These are highlighted in figure A) below.
The question also tells you that each turn of the alpha helix is 5.4 angstroms. One turn if shown in figure B) below.
Looking at the picture, it seems that each helix is about 7 turns long. As each turn is 5.4 A, the length of the helices must be approximately 7 * 5.4 A which is 37.8 A.
Therefore, using the information that the membrane must be about as thick as the helices in the transmembrane proteins are long, you can deduce that the membrane must be up to 37.8 A thick, however the membrane may not actually be this thick as the question from the paper doesn't specify how much of the alpha helices are embedded in the membrane (60%, 80%, 100%?). I guess the assumption is that the entire alpha helix section of the transmembrane protein is fully embedded within the membrane.

Are the alpha helices always straight within a membrane, or could they be bent?

From what I understand, alpha helices tend to be straight due to the strong hydrogen bonding which takes place throughout the structure, so it would be unusual for kinks to appear. However, there are structures called helix-turn-helix (common in DNA binding proteins) which form an alpha helix with a kink in it. There’s some good information on this here: https://www.sciencedirect.com/topics/biochemistry-genetics-and-molecular-biology/alpha-helix and here: https://www.sciencedirect.com/science/article/pii/S0005273618300518

Please do not answer questions that do not respect the model of this site.

From How do I write a good answer? in the [help], Answer well-asked questions. Not all questions can or should be answered here.

Thanks a lot for the answer. And yeah, the question asked is very vague. I had problems because of this vagueness.


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Why are certain cells more likely to be attacked in autoimmune conditions?
For example, type 1 diabetes occurs when the immune system attacks and kills beta cells in the pancreas. This is relatively common.
What makes beta cells more likely to be targeted than, say, alpha cells in the pancreas?

Autoimmune reactions each have their target antigens. They'll "attack" wherever those antigens are expressed. In the case of Type 1 diabetes, many specific antigens have been identified, and they all correspond to regions on the surface of pancreatic islet β cells. Ultimately, however, the immune reaction is targeting proinsulin and its processed product insulin, which are produced by solely by β cells. This is an interesting review covering the topic.


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Comparing angles between population vectors of neural activity
I am reading this paper on fear conditioning, where the following is given:

The n-dimensional population vector (activity of n neurons) evoked by the conditioned stimulus (CS+, auditory tone) before conditioning
The n-dimensional population vector (activity of n neurons) for the CS+ after conditioning

The authors write that: "[After conditioning] the CS+ population vector rotated out of the plane defined by the US [unconditioned stimulus] and the initial CS+". They conclude that this out-of-plane rotation corresponds to "new learning".
My question: Why is out-of-plane-rotation of neural activity vectors equated with a new representation/new learning?

They are not writing about "after conditioning" as you write, but, importantly, after extinction.
What they observe is that, during learning, the response to the conditioned stimulus (CS+) becomes more similar to the response to the US. In the n-dimensional space in which a population response lives, you can describe this as the population vector evoked by CS+ pointing closer to same direction as the population vector from the US.
We expect that during extinction, the CS+ vector will become different from the US compared to at the peak of conditioning. There are two general possibilities for how that could happen. The first possibility (perhaps the simplest one to expect) is that with extinction the CS+ vector returns to where it was initially, before the conditioned stimulus was paired with the US. You could describe this as merely forgetting what has been learned: responses to the US and CS go back to how they were before conditioning and it's as if nothing was ever learned.
However, that's not what these authors observe:

During within-session extinction, the CS+ representation did not revert and gained no more similarity to its initial representation before learning

Although they do observe that with extinction the CS+ vector moves away from the US, it doesn't move back towards the old response, it points in a new direction that is further both from the US response and from the original CS response. That can be described as an "out of plane" rotation, not towards either of the other vectors.
In case it's hard to think about these things in N-dimensional space, we can think of a more familiar situation in our own 3-dimensional world.
Let's say the US causes a response corresponding to "12 o'clock". The conditioned stimulus causes a response corresponding to "3 o'clock" before any training. As you do some conditioning training, pairing the US and CS, the CS response moves towards "1 o'clock", beginning to resemble the US response. During extinction, we might expect the CS response to move back towards "3 o'clock", maybe we'll find it at "2 o'clock". This would represent rotation in the "plane of the CS and US" made by the clock. Instead, the hand points out away from the wall, so it is further from both the original US at 12 and original CS at 3. That's the "new learning".

Thanks for the fast response! Right - extinction in particular. I think my point is a bit deeper: I understand that extinction is considered new learning, and hence the representation of the CS+ should not convert to its original form. I still don't see, however, why out-of-plane-rotation is "new learning". My understanding would be that previously inactive neurons would display activity now, as you describe the rotation would be into a new plane not defined by the original orthonormal vectors. Right? This seems to me a mathematical property, however, and very removed from behavioral learning?

..2/2 like, how do we know this mathematical out-of-plane-rotation is in any way physiologically meaningfull? What is the typical argument made for such an interpretation?

@Pugl Maybe you're thinking about it too deeply. It's just a geometric interpretation for the extincted CS+ response not returning to the original CS+ response. "New learning" just means that the CS, post-extinction, produces a response that is similar neither to the US nor to the original CS. That's it. It's evidence against a hypothesis that extinction means the response returns to how it was originally before learning.

@Pugl Another way to say it (sorry if I'm getting exhaustive) is that during learning of the CS-US association, the population response stays in the plane of the clock. It moves really just in one dimension defined by the plane of the initial CS and US responses, like the minute hand on a clock. Extinction moves the response in a different direction to that plane, into new territory not observed during the initial learning process.

Not at all. I appreciate all input. I actually would like to understand that better: Previously, they described in-plane rotation based on newly active cells. How is cell activity different between in-plane and out-of-plane rotations?

@Pugl Not sure what you mean by "they described in-plane rotation based on newly active cells". Might be worth another question. Also important that you have some grasp on what these vectors and planes are. Easiest to think about is with just 3 neurons, the population vector would be given by the activity of those three neurons on the X, Y, and Z dimensions. So if only X is active, you'd have a vector right along the X-axis. If only X and Y are active, you'd have vector pointing somewhere along the XY plane, but with no magnitude in the Z direction. Etc.


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Check simple human fetus answers
Based on this image and questions, could someone please check my answers?
Question 1: At which one of the following points will the blood passing the point be more oxygenated after birth than before it?
I would think at point P.
Question 2: At which one of the following points will the blood passing the point be significantly less oxygenated after birth than before it?
I think point L, because the blood should not lose any more oxygen as it travels through veins and the heart to the lungs.

Please see this sites policy for "homework" questions. Among other things, such questions must be appropriately tagged. Thanks!

It was not a homework question, but a practice question to demonstrate how a platform works before I take an online quiz, for which I have already included my thought process instead of asking for the answer outright. 2) I am unsure with which keywords to tag this as I am new to the community?

Please review the criteria on the page I listed. 2) homework is a tag, as is described on the that same page. In addition, please take the [tour] and to read [help] pages such as [ask]. The [meta] pages also contain many questions and answers about the expectations this community has for users. Please also consider whether [MedicalSciences.SE] is a better fit, but please do not crosspost, instead request migration. Also note that that site also has stringent expectations about prior research.

Well I added that tag, and I made a few changes since my responses would not have made sense.

Thank you for all of the downvotes, what a welcoming community. I thought it was just computer scientists with a toxicity problem.

I'm sorry you've found your first experience here unpleasant and I do understand that being downvoted can feel like a personal attack. However, downvotes are merely an evaluation of how appropriate a particular question is for this site. Your choice to view  that as "toxic" leads me to strongly encourage you to familiarize yourself with this sites [help] before your next post. Good luck!

@tyersome Sorry, I should have perhaps not worded that so harshly. Basically, where can I ask these style of questions to not upset people here? It was not even a homework question, but an example question for the official exam.

For question 1 I would agree with "P"
For question 2 the answer would be "M" although an argument could be made for "O". Consider that before birth "M" should be the points were blood contains the most oxygen since it is coming directly form the oxygen exchange While after birth it contains the least amount of oxygen becasue that exchange is lost and it instead only receiving blood from the body where most of the oxygen has been extracted. So the change at that point is about as large a change as possible.  I don't know how you got "L" it should have little to no change in oxygen content as both before and after birth it is receiving low oxygen blood from the body.

My point of view was L, M, and O are all equally deoxygenated after it goes through the capillaries at the very bottom of the image and should remain the same until the reach the lungs after birth. However, since the question focuses on AFTER birth then I agree it must be M because it has the biggest change from oxygenated to deoxygenated when the mother's maternal artery is disconnected.


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What is the Importance of Acid Alcohol and Sodium Bicarbonate during staining?
I've just learnt about staining during a histology class and I'm confused.
Why do we have to add acid alcohol and sodium bicarbonate to the tissue during Hematoxylin & Eosin staining? Is there any difference if we don't add them at all and just skipped to eosin instead?

Mind sharing the exact staining process that you are using?  There are slight differences and honestly I have no idea what sodium bicarbonate would be for, unless it's an attempt to get some ions back into soft water used for the experiment, but study has shown that to be a very ineffective way to go about it.

This is described with better detail (and better prose) in John Baker's classic description at StainsFile, but to summarize:  The procedure you're describing is a regressive hematoxylin stain with Harris hematoxylin.  To start off with, let's bear in mind that hematoxylin, a compound from the wood of the logwood tree, is actually used as hematein, by oxidizing it with sodium iodate.  (The colorless hematoxylin would naturally "ripen", oxidize over 4-10 weeks in the presence of air, even without that step, but this is more controllable)  Additionally, aluminum ions are added as a mordant, creating hemalum by binding =O and -OH on the hematein.  The Al atom is what actually binds to something in the nucleus (I found reference to lysine side chains of histone proteins and the phosphate of the DNA - I haven't sorted this part out yet).
When acid is added, it competes with aluminum, which can be thought of as Al3+.  The real bonding may be more covalent than that, but that point is considered controversial.  For whatever reasons, the H+ interferes with the Al3+ mediated attachment of stain to tissue, especially at sites of background attachment (differentiation).  The hematoxylin washes away somewhat, leaving lighter, more specific staining.  (this tactic is called a regressive stain, and is certainly not needed to do the procedure.  This is very much more art than science at this point...)
But ... the acidic hematoxylin has a reddish color, and it isn't as blue as it can be unless it is neutralized to around pH 8 - hence the second step with bicarbonate, though often plain tap water can be used.  That is an ordinary instance of a dye changing color with pH, and can be called "bluing the hematoxylin".

I'm just going to answer for the acid alcohol since I'm unaware of any purpose for using a sodium bicarbonate solution, unless you're referring to its use as a component in the eosin dye.

The acid alcohol is used between the hematoxylin staining and the eosin staining.  It's purpose is essentially to remove any non-specific hematoxylin stain (reference). Hematoxylin is acidophilic (hence its capacity to bind nuclear components such as ribonucleic acids & deoxyribonucleic acids), so if you wash the slide with a weak acid, the hematoxylin which is just "sitting around" unbound to an acidic cellular component will be washed away, making the colouration much cleaner since only the acidophilic cellular componenets will still have the blue stain on them.
On a more technical tack, it also keeps you from getting large quantities of hematoxylin in your eosin stain, thus allowing you to use your eosin stain for more slides before it no longer stains as well.


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Is there any problem with using a single stop codon with a single CDS in prokaryotes?
All protein coding sequences in the iGEM Registry are supposed to end with a double stop codon. Presumably, this is to decrease the potential for read-through, which could be problematic if one is putting together a polycistronic design.
If the design is intended to have only one CDS, however, and is targeted at a prokaryote (which have "backup" ribosome release mechanisms), then is there any problem with using only a single stop codon?

The impact of any read-through from a leaky stop codon in an expression unit with only one CDS would probably depend on a few things, mainly (i) where is the next in-frame stop codon, (ii) what are you trying to express, and (iii) how leaky is the stop codon?
In cases where the next in-frame stop codon is only a few base pairs away, there would probably be little impact, however in other cases the next stop codon could be far away. In these cases, there are two things which may cause an impact.
The first is that a long peptide sequence could be added to your protein, which depending on what you are expressing, may cause your protein to misfold or lose functionality.
The second is that you could get ribosome stalling, especially if any of the codons between your stop codon and a second stop codon require rare tRNAs. As you mentioned in your question, there are mechanisms for rescue in these scenarios, however if your CDS is expressed under a strong promoter on a high copy number plasmid, this mechanism may need to be mounted much more often than usual and cause burden on the cell. I should note that this is speculation on my part as I can’t find any studies which have shown this.
The actual impact any of these scenarios may have on your system would likely be strongly dependent on how leaky the stop codon actually is. As there are many examples of constructs which use only a singe TAA in their design with no apparent negative affects, presumably under ‘normal’ circumstances there is little risk to using a single stop codon.

I found an estimate of read-through in at least one gene being approximately 4%, which seems pretty high! https://www.ncbi.nlm.nih.gov/pmc/articles/PMC5133446/

If there is only one CDS, presumably there will be a transcriptional termination directly after the CDS (after the stop codon actually), which makes any read-through coming from using a single stop codon almost negligible.

Is there any issue with ribosomes getting stuck on the terminator?

Actually, somethings we tend to forget in prokaryotes are: 1. in general there are many ribosomes translating at the same time, the same mRNA. 2. Ribosomes start to translate directly on the nascent mRNA, as RNAP is transcribing.

What about on the other side? Can they get stuck on the end?

Perhaps I could add for eukaryotic systems which terminate all stop codons by a single protein, eRF1, this study by Schmied et al., showed readthrough of all stop codons to be less than 0.2% by wild-type eRF1, which seems acceptably low! I am unsure how this would transfer to prokaryotic systems which diverges in its use of two release factors - RF1 and RF2. I would of thought termination by UAA which is shared by both factors would be quite stringent.


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What determines walking direction in humans?
I've wondered about this for a while, without being able to come up with an answer.
Is there a body part that single-handedly determines the direction we're walking in? Is it a group of body parts that make us "steer" when walking?
I took a look at the behavior of some body parts while walking, and I came to the conclusion that:

it's not the direction the feet are pointed to, as you can turn them in or out while maintaining your path
it's not your head, of course, as you can be looking at any direction while maintaining your path
it's not your hips either, as you can move them freely without changing path (although I wouldn't suggest trying that in front of people)

As much as this sounds like a silly question, I still haven't been able to get to an answer. Any input is appreciated.

the brain which coordinates movements determine directions. You will note you can move in any direction regardless of which way you are facing. Whichever direction your feet are producing the largest net force is the direction you will travel.

Are you asking what part of the body controls voluntary motion?

It's the center of mass of your whole body, projected onto the floor, in comparison to the position of your feet; the center of mass will fall away from your feet, especially from the line that can be drawn between your 2 feet. So walking can be seen as continuous falling.

@KaPy3141: No, or more precisely, not necessarily.  Though what you describe is an efficient way of walking, it's not the only way.  Consider side-stepping (familiar if you're a cross-country skiier), walking backwards, or at a slow pace...

I Agree with @John, the feet that is in the floor generate a force that is going to move your center of mass. The other leg movement is also important because wherever it poses is going to determine the displacement of that step. Of course the rest of the body should move accordingly to achieve balance.

@james: Can't side-stepping be seen as standing on one foot, which pushes you in the direction of your center of mass?

@KaPy3141: Depends on how you do it, of course, and easier to do than describe, but basically you stand on the downhill foot, extend the uphill foot while remaining balanced on the other until the uphill foot is firmly planted, then transfer weight to the uphill foot, and lift the downhill foot to place it close to the other.


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Are there any animals whose genome haven't been sequenced yet?
I referred to a few websites and it seems like almost every animal's genome has been sequenced. However, are there any animals that haven't yet been sequenced? If so, can you provide the list here or provide the link to the website which provides that information??

There are 60,000 species of beetles, you really think people have sequenced them all?

@swbarnes2 We're on our way to doing so...

Wikipedia maintains an (incomplete) list of sequenced animal genomes here.
There are a few million living species of animals, we're not that close to sequencing them all, and listing them all doesn't make much sense.
I'd recommend starting from the animals that have been sequenced, and in particular, animals from particular families or smaller taxa, if you'd like more detailed information.

I see there are a lot of animals that we haven't yet sequenced but I mean are there like any animals that are beneficial to humans in some way, hence it's worth sequencing, but it hasn't yet been done? I thought about choosing chimpanzee but it has already been sequenced.

@ZetaFox "Beneficial to humans in some way" is a pretty broad criterion that could likely include every animal there is with some justification.

@ZetaFox A lot of focus is actually on those that are harmful to humans, which is why you'll find thousands upon thousands of sequenced strains for many pathogens.

@jakebeal I kind of read OPs comment as "beneficial to sequence" but that's a good point to emphasize in case I was reading too much into it.


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Can a cell receive multiple copies of an insert when using different MOIs?
I want to transduce a cell line with virus that carries a specific insert.
When using different Multiplicity of Infection (MOI), I expect to get different percentage of transduced cells, but is it possible to get more than one insert integrated in the cell genome?
I use high MOI and I always get 1 copy per cell, so I am not sure if increasing it a lot may also affect the number of in integrated inserts.

Yes, if you use a high enough MOI, you'll start to get double infections. Usually, anything >1 MOI will guarantee a high chance of double infections. If you're using a high MOI and only seeing single inserts, it might be due to your cells being difficult to transfect. If you look at this site: https://manuals.cellecta.com/lentiviral-construct-packaging-and-transduction/v10a/en/topic/lentiviral-titer-calculation it should give you a general reference for what % double transfected cells you should see. The difference between transfected cells and MOI % should estimate double infections assuming 100% efficiency.
Some cells are less efficient at being transfected, and it's a good idea to get a functional titer on your target cell line which is very easy if you have a fluorophore in the plasmid you're trying to insert.
I reread what you're getting at, and it's generally accepted that MOI under 0.4 won't give you double transfections.


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Calcium ions and bacterial transformation
What is the mechanism by which calcium ions and heat treatment allow the bacterial membrane to become permeable, allowing the uptake of plasmids?

DNA has an overall negative charge. The phosphate heads of the lipid molecules that make up the cell membrane, along with the LPS molecules on the outer membrane, are also negatively charged. This means the plasmid is normally repelled by the cell membrane.
What Ca2+ ions do is shield the negative charges on each side, making the overall situation electrostatically neutral.
This site has a nice animatic for this process: http://www.dnai.org/b/index.html

Thanks owlofathena

There are two things that need to be corrected in your answer. The first is that the effect of calcium is on the bacterial cell wall, not the cell membrane. The second is that "lipid" should be replaced with the specific membrane constituent of Gram-negative bacteria — lipopolysaccharide. There is a short Wikipedia entry for calcium chloride transformation.

You're right that I should have included LPS along with phosphate groups as contributing to the negative charge on the cell surface. That was sloppy of me. Edited. Thanks!


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Pipetting: do human experimenters need liquid class information?
I've been working on a protocol standardization project where, among other things, we want protocols to be able to be run equivalently by both humans and robots.
Something that I've noticed in doing so is that there's a radically different amount of information typically assumed needed by a human vs. a robot when it comes to pipetting.
With a robot, it is generally recommended to understand the nature of the liquid thoroughly, specify the liquid class (which includes information on viscosity, volatility, polarity, etc), and do quite a bit of quality control to make sure than when you want to pipette 100 µL you actually get 100 µL. A typical example is shown on this page by Hamilton.
With a human experimenter, however, the protocol pretty much always just says "pipette 100 µL" and assumes that the human can figure out all of the potential issues that we have to be so careful for with robots.
Is this actually a safe assumption, or just something that we don't typically check during protocol development for humans? Is it really intuitively obvious for nearly all liquids, or should our protocols be providing more guidance for humans as well?

I have answered this question from my viewpoint as a student, but I think it could benefit from an answer(s) from someone who is on other side of the "experience" divide, and is more used to giving advice than taking it.

I don't wish to be overtly critical, but how is this a question about biology? I have already removed the incorrect tag "synthetic biology", but even as a methods question it seems to be only answerable in a subjective sense as human psychology, or in a legal sense in relation to the laws of a particular jurisdiction  relating to some form of industrial safety.

@David I'm really not sure how you're coming to that conclusion: this is a problem of biological protocol replicability.

@David I've added the lab-techniques tag as well; if you disagree with this question, then what do you find different from other lab-techniques questions on this site?

Lab techniques questions are of the form "How do I (best) do such and such?" That sort of question allows an objective answer "You can do it by..." Your question seems to be of the form whether it is necessary to write instructions to people performing a particular (common) laboratory technique. This can only be answered subjectively. I have voted to close this question on that basis, i.e. it is a matter of opinion.

@David Interesting; my own assessment, which supported my choice to ask, is that this falls solidly on the good subjective side of the bar, but of course that is, itself, a subjective judgement.

For what it's worth, I find this a particularly interesting question which already has interesting answers and a very refreshing change from most questions we get. One of the long term problems we've had here is the relative dearth of professional biologists asking professional-level questions so I am really surprised to see this already has two close votes! I would have thought this is exactly the sort of question we want to attract and support.

"the human can figure out all of the potential issues" Yes, it's one of the first things we got taught in chem lab. You could start with the information provided in those courses.

In my experience it is very rare to see a protocol for humans that describes how to pipette a certain liquid, but I don't have as many years pipetting as others do.
In general, it is left to the experimenter to observe the behavior of the liquid and act accordingly. It is quite easy to determine when a liquid is leaking (e.g. ethanol) or when it is very viscous (e.g. glycerol), and determine that a change in technique is required. In this case, I would ask someone else in the lab how to deal with the situation. This is something that robots cannot do.
However, not everyone has expert help available, and although it is fairly obvious when a technique isn't working, often the solution is not intuitive. Additionally, the advice of others in the lab could be unhelpful or even inaccurate. For example, I was told to cut the tip off a pipette in order to pipette Tween-20 more easily, however in reading for this answer have discovered that this is not recommended at all! (See eppendorf FAQ linked below).
Therefore, I would say that, unless the liquid is to be pipetted in the same way as water, it is always useful to provide information for those who are not knowledgeable or do not have easy access to expert practitioners.
For some examples of the sorts of problems that experimenters come up against, and therefore which liquids especially could use instructions in protocols, see this set of FAQs from eppendorf on handling liquids.

Wow... I think it's particularly revealing how answering this question actually ended up causing you to discover that an "oral tradition" answer was incorrect! It seems that pipetting actually has a lot of knowledge required in order to do it correctly?

As with so many things we do in the lab, we are often in a rather high local minima of 'what worked for postdoc Bob 15 years ago and has been handed down ever since'

Many don't know that pipettes are calibrated for water solutions. Organic solvents (like ethanol) have a different partial pressure and can't be measured with a water calibrated pipette.  This is crucial in some experiments.

The most finicky pipetting situations I've encountered are with regard to emulsions. The worst example I've encountered is the 10x Chromium, which asks you to dispense 95uL on a P-200 multichannel over the course of 30s to avoid breaking emulsions.  But plenty of other emulsion-based protocols (often single-cell-related) specify slow pipetting

I work on software that controls one model of robotic pipetter.  There are a few reasons why we use complex liquid class definitions:

One of the selling points of using a robot is high speed.  You can handle a wide range of viscosities if you set a conservative (slow) speed for aspiration/dispense, but for solutions that don't require this, we'd like to be able to move more quickly.  In many cases, the different liquid classes are about optimizing the time required while still maintaining the desired precision and accuracy.

Robotic pipetters will often dispense multiple times after a single aspiration of a larger volume.  To do this accurately, we compensate for a variety of effects, including the effect of the column pressure, which depends on the reagent density.  In the end, this is also largely about improving speed.

In some applications, precision is especially important and you can use settings for the robot that improve precision.  Let's say you have something that is extremely sensitive to getting the right pH.  You could pre-titrate a buffer to the correct pH and load that into the system, but if you're trying to produce an assay grid on a microplate, you'd have to make a bunch of buffers.  A well-calibrated robotic system can mix from stock buffers to produce the assay directly, since it has the necessary precision for mixing very small volumes.

Not having much personal lab experience since school, I can't speak to how precisely/accurately pipetting is done in labs by human beings.  It probably varies quite a bit from lab to lab and likely depends on the skill and tiredness level of the person doing the work.  I did it enough times to know that even basic pipetting of aqueous solutions has a bit of a knack to it and was glad when I was done with the labs that required it.  In any case, I thought it might be useful to have an idea of why the robotic control seems so much more complicated; it's mostly because robots are trying to do it faster and without the benefit of being able to observe the reagent's behavior while using it.

I appreciate you sharing your expertise about the robotics, but that's the side that I'm more familiar with as well. I'm afraid however, that as currently written this answer isn't shedding terribly much light for me on the question about whether humans need more information than they are typically given.

@jakebeal, it doesn’t quite say it explicitly, but I interpreted this answer as saying “humans don’t pipette in the quite the same way, and therefore may not need as many details.”

@MattKrause Agreed; I was hoping for a little bit more description and basis for that conclusion.

@jakebeal I'm not aware of nay research on the topic, but I would suspect that even if you give explicit instructions to human operators, they might try to apply them, but most likely they'll largely fall back on their learned muscle memory and habits from their training/practice.  In a sense, I'd say the humans already have the liquid class information in their brains, to the extent that they've encountered those types before.  Humans also have the major benefit of being able to look at the pipette tip and see if there's still fluid stuck inside and can adapt accordingly.

Just to be clear, it has been many years since I have done real lab work, so I am certainly not an expert on the specific question.   However, I have a fair amount of experience writing clear instructions to cover complex (and often bespoke) processes.
As Dan Bryant has explained, robotic systems absolutely need the additional detail in order to complete their tasks optimized for speed (and a control method without the built in feedback loops that come with a human).
My experience writing procedures for humans is that same level of detail will actually be a significant negative, at least on techniques that are considered standard knowledge.   Providing what humans consider 'unnecessary detail' tends to result in a problem where your procedure is 'skimmed' for key information, which may result in lower adherence rather than improved results.
Another consideration is your level of expertise - do you know every situation where a task specific pipette is required?  Because if you specify it once, you create an expectation that you will specify it every time, so any situation you miss may cause an error, and the probability of error is higher with operators having less experience - the primary audience for detailed procedures.
There are solutions, but they tend to depend on your documentation standards and the documentation tools in use.   The simplest is often to move the robot only detail into an appendix (or something similar, such as a collapsing box).  This means the person programming the robot will be able to drag the detail out when required, but the 'visual noise' for the humans will be reduced.

Re: specifying robot details, most vendors will ship liquid class information with the tool covering the majority of cases, so you generally only need to go through a careful calibration process on your own if you need particularly tight performance or you have an unusual fluid.  The parameters and configuration process will also vary between vendors/tools, since, among other things, there are a variety of different technologies in use that all operate slightly differently.  It's worth specifying the liquid class you used for replication, but would be hard to distill to tool-independent info.

That is a very good point about including unnecessary detail. If this information is included it should be in a subtle/optional way so that it won't distract people who don't need it.

In my experience, I've never seen more than "pipette 100 μl" or at most "caution viscous liquid". Also I've only been taught how to pipette in a general manner and never about how to do it with special liquids except for the "cut the tip off" advise so before Noah's answer I had never heard of reverse pipetting or using a multivette to accurately pipette glycerol for example as described on the eppendorf website. However, when pipetting glycerol for example I could tell that I needed to pipette it slowly and I could check visually that I had (about) the right volume using the marks on the tip. So compared to a robot I can use my eyes to adjust and it's good enough in my particular case and with a single tube it is not optimal and it may not hold true with a multichannel pipette so having good and better pipetting habits would be better for reproducibility.
Now if I can see the "cut the tip" advice and the standalone "pipette 100 μl " instruction work to some extend, "commercial" grade protocol would probably have the instruction written as "using a multivette (or using the reverse pipetting technique) pipette 100 μl".
Another thing to consider is the definition of protocol which if we take wikipedia's definition is not just a succession of steps to perform :

In addition to detailed procedures, equipment, and instruments, protocols will also contain study objectives, reasoning for experimental design, reasoning for chosen sample sizes, safety precautions, and how results were calculated and reported, including statistical analysis and any rules for predefining and documenting excluded data to avoid bias.

What we usually consider as a protocol is the "Quick protocol" with only the steps to perform where you're more likely to find the "pipette 100 μl" type instruction. But if you consider the more detailed protocol you'll find more information. I don't have an example for liquid handling but if you consider the qiagen miniprep protocol depending if you take the quick version or the full one you may not be doing the same thing. Let's take step 2 as an example :

quick protocol :

Add 250 μl Buffer P2 and mix thoroughly by inverting the tube
4–6 times until the solution becomes clear. Do not allow the lysis reaction to
proceed for more than 5 min. If using LyseBlue reagent, the solution will turn
blue.

handbook :

Add 250 μl Buffer P2 and mix thoroughly by inverting the tube 4–6 times.
Mix gently by inverting the tube. Do not vortex, because this will result in shearing of
genomic DNA and contamination of plasmid. If continue inverting the tube until the
solution becomes viscous and slightly clear. Do not allow the lysis reaction to proceed for
more than 5 min.
If LyseBlue has been added to Buffer P1, the cell suspension will turn blue after addition
of Buffer P2. Mixing should result in a homogeneously colored suspension. If the
suspension contains localized colorless regions, or if brownish cell clumps are still visible,
continue mixing the solution until a homogeneously colored suspension is achieved.

If we consider only the mix part here, "gently" is the key word. If you've been handed only the quick protocol and you're an inexperience user even just by inverting too strongly you may end up shearing the genomic DNA and thus contaminate your sample what can impact your downstream application.
So the bottom line is that "pipette 100 μl" may be enough for experience users and/or some applications on a quick protocol but you may also want to have the instruction "pipette 100 μl using a multivette (or using the reverse pipetting technique)" on a more detailed protocol for inexperience user or troubleshooting purposes.

Thank you for the information about the miniprep protocol! I'd always assumed that thoroughly meant that it didn't need to be/shouldn't be gentle. I normally shake the tube viciously for that step...


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Confusion regarding the Kir2.1 inward-rectifying potassium channel
I was trying to find out more about gustation and the transduction of sourness when I came across the supposed inward-rectifying potassium channel $\ce{K+_{ir} 2.1}$. Here's the thing, despite being inward rectifying, all the articles I read about it involves it sending potassium ions out of the cell. I thought that maybe there is a special mechanism that makes it outward-rectifying, or just switches off its mechanism for inward rectification but to no avail.
So here's the context: On taste buds that detect $\ce{H+}$ ions, when intracellular pH rises due to intake of protons, potassium channels will be closed which causes accumulation of $\ce{K+}$ and thus depolarisation. The article pinpoints $\ce{K+_{ir} 2.1}$ as one of the potassium channels involved, saying that its inhibition will allow for depolarisation. This doesn't make sense to me, if this channel is supposed to prefer intake of potassium ions and almost entirely prohibit outward flow. If we want to accumulate potassium in the cell, shouldn't this channel be left open?
The only thing I can think of is that this channel remains open at membrane potentials near that of resting potential. So maybe then there is no selectivity over which direction the ions travel, so there is a net outward flow?
I'll link some of the articles I mentioned below:
The K+ channel KIR2.1 functions in tandem with proton influx to mediate sour taste transduction
Inhibition of Kir2.1 channel-induced depolarization promotes cell biological activity and differentiation by modulating autophagy in late endothelial progenitor cells.

Short answer
"Inward rectifying" doesn't mean "only allows inward current". Current through inward-rectifying channels can be (and often is) entirely outward in physiological settings.
Explanation
For a non-rectifying channel, there is a linear current-voltage relationship. If you plot current through the channel at various holding voltages, you get a straight line.
For inward-rectifying potassium channels, you get less current as you depolarize, like this:

(from Wikipedia)
At negative voltages, the channel is pretty linear and strongly inward (negative). However, at more positive voltages, the linear trend is attenuated.
However, this relationship does not upset the equilibrium potential for a given ion. For a potassium channel, if you have a cell with a potassium reversal potential that is more negative than the membrane potential, you'll still get an outward flow of potassium ions (in the figure, note all the lines above 0, representing voltages more positive than the potassium reversal potential; they're positive/outward even if they're small). What is different for the inward-rectifying channel is that as you depolarize the cell, the current does not increase the way you would expect it to increase for a non-rectifying channel.
For example, for a non-rectifying channel, you'd expect with a potassium reversal of -90mV, the current would be the same magnitude but opposite sign at -120 mV and -60 mV (30 mV above and below reversal). For a inward rectifying channel, you will still have the same change in sign, but the magnitude will be less at -60 mV than at -120 mV.

its inhibition will allow for depolarisation

Any channel permeable to some ion (or combination of ions) will tend to move the voltage towards the reversal potential for that ion/ions. So, as long as potassium reversal is more negative than threshold, all potassium-only conductances are inhibitory, whether they rectify or not.


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Surface Plasmon Resonance (SPR) Predicting Molecular Weight Limit?
Just asking, can we predict how low molecular weight (LMW) analyte our SPR system can detect based on our LOD (limit of detection) and our sensitivity on another protein detection? So right now our developed system can detect alpha-synuclein (14 kDa) with LOD around 5fg/ml. Can we predict how LMW can we detect? Is there any reference paper or something. Because as far as my knowledge we can get the change of RII based on the experiments first. But right now my case is I want to test the limit of our SPR devices to detect LMW (I'm choosing what LMW analytes to buy right now)
Thank you! any discussion or comments are very appreciated

Can you please expand your acronyms?

@jakebeal sure!


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Do carboxylesterases, arylesterases and acylesterases count to the enzyme class of lipases?
I am currently examining the acyl transfer catalysis activity of several enzymes that I was told are all lipases. Through reading a couple of papers including these enzymes I found out that all of them are hydrolases with promiscuous acyltransfer activity. Specifically one of the enzymes is a carboxyl-esterase, one is an aryl-esterase and the third one is an acyl-esterase. I know that esterases are part of hydrolases and lipases are basically fat-splitting esterases. Lipases also catalyze transesterification = acyl transfer, just like all of the enzymes I am working with do. I just don't understand the classification of all these enzymes as lipases. Can someone explain the connection to me?

Could you clarify. You explain that you understand that the enzymes in question are all "fat-splitting" i.e. lipid-hydrolysing esterases. More specifically they hydrolyse lipids that belong to the class ester (e.g. triglycerides) rather than lipids in general such as free fatty acids or cholesterol. Such enzymes are given the trivial generic name lipase. (They have more specific systematic names.) What is it that you do not understand? Do you want examples of systematic names or what?

The assay we are working with is using para-nitrophenyl butyrate as a substrate. This substrate is usually used for investigating esterase, as well as lipase activity. I read that "Lipases can be distinguished from carboxyl esterases by their substrate spectra, using p-nitrophenyl palmitate (cleaved by lipases) versus p-nitrophenyl butyrate (cleaved by esterases)" (Source: https://www.scielo.br/pdf/cta/v31n3/a09v31n3.pdf). So shouldn't the enzymes I am using be classified as esterases, rather than lipases?

I think your problem is one of the use of trivial names (such as lipase) and systematic EC names, which I have answered here Origin of enzyme names.

This is further to my previous comment which was autogenerated when I voted to close your question as already having an answer. I would stress that "lipase" is not a systematic name but a trivial name, that the names people use for enzymes (as of other things) depends on what they were taught and what interests them. If you are interested in lipids, the name lipase appeals, if you are a chemist, not. See also my answers for other enzymes here and here.