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What type of spider is this please?
Can anyone identify this spider please. Found in Herefordshire UK

Welcome to Biology.SE! Identifications questions should include: 1) one or more clear closeup photos of the organism (photos from multiple angles if possible); 2) habitat information; 3) an estimate of the size of the organism; and 4) any behaviors that you observed. Please [edit] your post to include all this essential information. ——— Please also take the [tour] and then go through the [help] pages starting with [Ask] questions effectively on this site. Thanks!

It looks to me like a young Steatoda of some sort, but that's really just a guess based on the shape and posture of the brown banded legs and the shape and relative sizes of the cephalothorax and abdomen.  The photo is good, but it would help to have some sort of size comparison like a coin, plus perhaps a bit more flash to expose any patterns on the top surface of the body.


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Safety in Liposomal Amphotericin B
From reading about Amphotericin B antifungal, I understand it is quite a toxic medicine for kidney, but I am not able to find any details as to how Liposomal version of same medication is so safe, in end are not we eating same chemical, how doe s liposomal version become less toxic?

This is an interesting question.  Initially, the mechanism for the reduced toxicity (1992) was unknown, as stated in the abstract of a well-cited paper.  A paper from Oxford Radcliffe hospital in 1998 said that 15.5% of all their spending on antimicrobial agents was on these new, expensive drugs, available in many formulations, and argued the evidence supporting their use was scanty with a key study available only as an abstract.  A review in 2013 seems more favorable, but I haven't looked at the full text - by that time, one of the products with reactions at the injection site had been withdrawn, and the costs were coming down.  At the level of the individual, these stories are poignant.
The paper from Oxford Radcliffe wrote of one formulation "It has been suggested that the large particle size of ABLC (1.6–11 μm) leads to its rapid clearance by reticuloendothelial cells, resulting in lower plasma levels."  This idea has been pursued in numerous other papers (example), but I lack the expertise to evaluate it against other potential explanations.

source: Stack Exchange

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

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larvae in my balcony
yesterday I found lots of larvae on my balcony floor.  I couldn't find out where they came from. I put some in a jar to search for them and today they started turning into pupae.
Can anyone tell me what the insect is and how to get rid of it?

Welcome to the site. Take a [tour] and visit the [help] for details on how to ask a good question. For identification questions we also need a location (to local area, don't need your street address), what's in the environment (e.g. nearby plants, animals etc) and preferably size or some sort of scale to refer to. As another point you are asking for an identification of a pupa - this is entomology (insects etc), not botany.

keep the pupae in a jar so we can identify the adult later

Oh, I know those things.  I've seen the empty pupa cases, and I've seen the larvae.  I did not know the pupa cases were from those particular larvae, so you've already found uot more than I knew!

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


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How much genes can be knocked down at the same time
When experimenting on mice, gene knockdown seems to be a useful technique to deactivate individual genes to study what they are doing. In practice what's the maximum of genes that can be knocked out at the same time?

I think this question needs clarifications / constraints. Assuming we're talking about mouse knockouts, are you interested in the technical limitations of creating knockouts, or are you asking about the biological limitations of KO mice (i.e. a minimal mouse genome)? Must that mouse live a full mouse life? Should that mouse be able to make more mice?

My interpretation was that he is asking about transient reduction of expression (e.g. antisense oligos) rather than genomic knockouts.

My confusion is the discrepancy between the title (knock down) and the text of the question (knock out), though I think my initial comment stands in either case.

As the OP asked about mice, I am mostly going to talk about organism-level things here.
First of all it is important to distinguish between knock-down and knock-out. Knock-out is where a gene is either removed or made non-functional and as such will never be able to be turned on again. This may in some cases be compensated for by having heterozygous animals, where one allele contains the normal gene and the other allele the non-functional, so that normal expression from the normal allele is enough to produce a normal animal. The heterozygous animals are then mated to produce homozygous (both alleles removed) knock-out progeny.
Knock-down on the other hand is either a transient reduction in expression (mostly done in cells, I don't think there are any methods for this that are viable at the whole mouse sort of level) or a conditional knock-out where expression of the gene is made conditional on a treatment, whereby you remove the treatment and the gene stops expressing. For example you can make a transgenic mouse with a tetracycline dependent promoter for expression (or not) of the transgene (Tet-On/Tet-Off). Knock-down in the strictest sense is only a reduction in the level of expression and may not be complete removal of the expression, which may result in viable cells through residual expression where a knock-out, with complete removal, might not.
However, back to the question at hand:
The answer to this is a rather unsurprising it depends on the genes being knocked down
Knockdown a single gene essential for cellular function such as something in a major metabolic pathway, and cells just won't function, so cells and animals will be non-viable. Knock-out or knock-down of non-essential genes and animals may survive, but also may not. As far as I know at this point in time we have no way of knowing whether knock-down or knock-out of something will result in viable progeny other than experimentation.

Why the downvote? - please let me know how I can improve my answer.


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Has this 2018 report of nematodes frozen for 30 to 40,000 years been confirmed? Has Carbon 14 dating confirmed their age?
Gizmodo's 2018 article Russian Scientists Claim to Have Resurrected 40,000-Year-Old Worms Buried in Ice says:

A team of Russian scientists is lining themselves up to be the opening cast of a John Carpenter film. Earlier this month, in the journal Doklady Biological Sciences (Viable Nematodes from Late Pleistocene Permafrost of the Kolyma River Lowland), they announced they had discovered ancient nematode worms that were able to resurrect themselves after spending at least 32,000 years buried in permafrost.
The discovery, if legitimate, would represent the longest-surviving return from the cold ever seen in a complex, multi-celled organism, dwarfing even the tardigrade.
[...]
It’s definitely not out of the question that these worms could have been revived after so long, according to Robin M. Giblin-Davis, a nematologist and acting director of the Fort Lauderdale Research and Education Center at the University of Florida.
“Theoretically, it is possible that if the organisms are protected from physical damage that would compromise their structural integrity during their frozen internment, they should be able to revive upon thawing/rehydration for very long periods of time,” he told Gizmodo via email.
At the same time, the team’s findings could still be a dud. “The biggest issue is the potential for contamination of ‘ancient samples’ with ‘contemporary’ organisms,” he said.
While the researchers do admit the possibility of contamination in the paper, they say it’s unlikely. They cited strict procedures to ensure complete sterility. And given that the ice samples were buried 100 feet and 15 feet down, respectively, they argue it’s implausible modern day nematodes could have wormed their way that deep.
The researchers identified some of the worms in the 32,000-year-old sample as belonging to the genus Panagrolaimus, and some of the worms in the 40,000-year-old one as part of the genus Plectus. Byron J. Adams, a nematologist at Brigham Young University who has studied nematode species capable of surviving extreme conditions, said the researchers’ claims seem credible, based on what we know about the biology of some modern-day nematodes.
“I’d love for it to be true,” Adams told Gizmodo. “We see what we think are prolonged stasis in Plectus and Panagrolaimus in Antarctica but have difficulty carbon dating them.”

and contains the image shown below.
Question: It's now been three years; has this July 2018 report of nematodes frozen for 30 to 40,000 years been confirmed? Has Carbon 14 dating confirmed their age? Since this period is many half-lives, there should be very little remaining Carbon 14 and and at least their approximate age should be straightforward to confirm and contamination with "contemporary organisms" fairly easy to rule out.

Selected images of the frozen worms said to be recovered from ancient permafrost samples dating back at least 32,000 years ago. Image: Shatilovich, et al. (Doklady Biological Sciences)

Background and relevance:
See Space SE's What precautions are planned to prevent samples returned from Mars crashing and releasing organisms on Earth?

Genesis crash site scenery; An earlier sample return mission broke open upon impact and exposed contents to the environment
This now has renewed interest because of another long-permafrost-preserved animal's ability to survive:

This animal survived 24,000 years frozen in the Siberian permafrost
Frozen in Siberian Permafrost for 24,000 Years, Microscopic Animal Comes Back to Life

“The takeaway is that a multicellular organism can be frozen and stored as such for thousands of years and then return back to life—a dream of many fiction writers,” said (Stas) Malavin. (a co-author of the study A living bdelloid rotifer from 24,000-year-old Arctic permafrost)

From Wikipedia's Bdelloidea:

The main characteristics that distinguish bdelloids from related groups of rotifers are exclusively parthenogenetic reproduction and the ability to survive in dry, harsh environments by entering a state of desiccation-induced dormancy (anhydrobiosis) at any life stage. They are often referred to as "ancient asexuals" due to their unique asexual history that spans back to over 25 million years ago through fossil evidence.

Asked in meta: How to tag questions about animals or other living things surviving being frozen for a long time? Is a new tag needed or just adjust cryonics tag?


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Detecting multiple compounds on a lateral flow immunoassay – is this a case of monoclonal vs polyclonal antibodies?
I am working on developing lateral flow immunoassays for drugs of abuse and needed some advice on detecting multiple drug compounds in one immunoassay.
As an example, an existing LFIA on the market tests for amphetamines, and detects the following compounds as positive at varying levels:

d-Amphetamine
d,l-Amphetamine
l-Amphetamine
3,4-Methylenedioxyamphetamine (MDA)
β-Phenylethylamine
Phenylpropanolamine
Tyramine
p-Hydroxynorephedrine
(±)-Phenylpropanolamine
p-Hydroxyamphetamine
d,l-Norephedrine

These compounds are all tested in one test strip — not multiplexed; the presence of any one of those compounds above the cut-off level will cause the single test strip to show a positive.
I don't quite understand how antibodies work and have a few theories:

Polyclonal antibodies have been used on the test line, which means all these compounds are detected
A mixture of monoclonal antibodies have been used, to detect all these compounds
These compounds are all chemically similar enough to be reliably detected with a single monoclonal antibody

Which of the above would be correct? Or am I misunderstanding this entirely and the answer is something different?

Background: I worked for a number of years as a product scientist for a fairly well-known antibody company, designing and producing polyclonal and monoclonal antibodies from several species.
Of your three theories, any one of them could be correct. Polyclonal antibodies are the cheapest, easiest, and generally fastest way of producing antibodies to a particular target, and I would imagine that they're used pretty extensively in LFIAs and other clinical immunological tests. However, polys have a major limitation: reproducibility. Animals' immune responses can change over time, even with regular booster immunizations, so it can sometimes be difficult to maintain the exact same specificity and sensitivity over multiple lots or batches, especially if you have to change animals. One way around this is to make very large lots, but they may end up degrading over time, even if kept at very low temperatures.
One way to get the recognition of multiple epitopes that polyclonals have with the epitope stability that monoclonals have is to make a "multiclonal" mix of monoclonal antibodies. This could be anywhere from 2 to a dozen or more clones, depending on the number of targets you're trying to analyze.
A single multiclonal antibody is also possible in this circumstance. While a monoclonal does only recognize one epitope, that epitope could be rather general and exist on multiple molecular configurations. An example of this type of antibody is Cell Signaling's Phospho-Akt Substrate (RXXS*/T*) rabbit monoclonal antibody (clone 110B7E). Instead of recognizing phosphorylated Akt/Protein Kinase B, this clone recognizes its phosphorylated substrates that have an Arg-X-X-pSer/pThr motif in them. It has been shown in tandem mass spec studies to bind to hundreds of unique peptides with this motif.
In the context of this particular question, it is perfectly reasonable to believe that the mixed amphetamines LFIA could rely on a single clone that recognizes a generalized "amphetamine-like" epitope that is present in all of the molecules listed, but (hopefully) not in other chemicals that are not drugs of abuse.

In general, monoclonal antibodies are used as primary antibodies and conjugated to the nanoparticles due to their ability to bind specifically to a single epitope of an antigen (1). Even a small change in the structure of the epitope can affect the binding to antigen, so this automatically makes your last statement impossible because a single monoclonal antibody can only recognize one epitope of the same antigen (2). In contrast, polyclonal antibodies are mostly used at the test band because they can recognize a host of antigenic epitopes, meaning that they are able to recognize highly similar compounds that vary in a single or small number of epitopes, making them ideal for generating signal for detection (1,2). But keep in mind that use of polyclonal antibody can also lead to increased risks of cross-reactivity (false positive result) (1,3).
Resources
(1) Qriouet Z, Cherrah Y, Sefrioui H, Qmichou Z. Monoclonal Antibodies Application in Lateral Flow Immunochromatographic Assays for Drugs of Abuse Detection. Molecules. 2021 Feb 18;26(4):1058. doi: 10.3390/molecules26041058. PMID: 33670468; PMCID: PMC7922373.
(2) Neil S. Lipman, Lynn R. Jackson, Laura J. Trudel, Frances Weis-Garcia, Monoclonal Versus Polyclonal Antibodies: Distinguishing Characteristics, Applications, and Information Resources, ILAR Journal, Volume 46, Issue 3, 2005, 258–268, https://doi.org/10.1093/ilar.46.3.258
(3) Reschly-Krasowski JM, Krasowski MD. A Difficult Challenge for the Clinical Laboratory: Accessing and Interpreting Manufacturer Cross-Reactivity Data for Immunoassays Used in Urine Drug Testing. Acad Pathol. 2018;5:2374289518811797. Published 2018 Nov 21. doi:10.1177/2374289518811797

This is not correct, especially your bolded statement. It is perfectly possible for a single monoclonal antibody to recognize multiple targets, even if they are composed of different amino acid sequences (in the case of protein targets) or atoms, in the case of small molecule targets. Any one of the OP's proposed mechanisms could be correct.

@MattDMo if this is the case, I would accept that comment as an answer – that there isn't enough information to tell which is the case.
Certainly would appreciate any other feedback to improve the question.


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How fast do ions move in body when they go in and out of the cell membrane?
Do we have any measurement tools to measure the speed of ions moving in a human body, specifically going in and out of the cell membrane? Do we have any tools to know where an ion is at a specific time?

Why do you want to know? Might https://meta.stackexchange.com/q/66377/401068 apply to your question? See also https://en.wikipedia.org/wiki/Brownian_motion

We do - check out patch-clamping


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Co-culture models of bacterial invasion: where to find them?
I am interested in models showing how pathogenic bacteria (namely Escherichia coli O157) can overcome commensal species (thus, causing a disease). This is a basic concept in biology but I can't find a proper article describing it.
I am looking, essentially, for a co-culture experiment with a pathogenic and a non-pathogenic bacteria that shows how the former can outcompete the latter. Is there a reference to such experiments?
Thanks

Are you sure you know what a commensal species is because the Wikipedia definition is incompatible with one member overcoming another: "Commensalism is a long-term biological interaction (symbiosis) in which members of one species gain benefits while those of the other species neither benefit nor are harmed" ? And which species do you maintain Escherichia coli O157 outcompetes — could you cite references on this point?

@David, I believe Gigiux is using commensal with respect to the hosting organism (e.g. human), contrasting those species with E. coli O157, which is not commensal with the host.

@mgkrebbs OK. But does the O157 strain have to complete with the commensal strain? For what? There are dozens of different bacteria in the gut managing to share resources.

@David, This usage of commensal is pretty consistent with the usage in most gut microbiome literature. Although I agree with you that it is probably used incorrectly most of the time, it seems like a fair enough label to apply until a specific harm or benefit for the host has been empirically demonstrated.

@MikeyC OK I misunderstood the OP with regard to commensal, but — and I am asking with skepticism but out of ignorance — does  O157 compete with K12 (or whatever) as his question implies? Because this is the crux of the viability of what he proposes. To do competitive growth experiments it would have to be established that the two strains complete for some nutrient, and what that nutrient is to be able to construct a growth medium. If this is true/know one would imagine it is also known what O157 does better — induce some enzyme, or whether its toxin affects K12. I think the OP should clarify.

I do not consider a request for a reference as question about biology and therefore will vote to close this. I found an answer by a simple internet search resulting from the comments above. Vet Sci 2018 5 93 https://www.mdpi.com/2306-7381/5/4/93#.

@David you are right: I meant commensal in terms of the interaction with the human host. I read already the article you indicated. But I am looking at a system describing how a bacterium pathogenic for human outgrows (I used outcompete because grows more than  the harmless species) one that (1) is not pathogenic and (2) that was initially present at a density higher than the pathogenic form.


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Why does this tree have a round hole?
One of my soccer students was hiding behind a tree when he discovered this hole. I've tried Googling it but I couldn't find out why this tree has a hole.

It looks like somebody just pulled out the outer layer of bark. I don't think there's anything special about it.

I agree. There was some sort of cut or other injury to the outer layers of the tree (bark and some of the living tissue just underneath it), and when the bark grew back it was not in concert with the rest of the bark, and formed a different pattern.


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What is the typical arrangement of amplified genes in archaea genome?
I have to check if some genes are amplified with bioinformatics tools, but, in order to do it, I need to know what this means from a biological point of view.
It seems not so difficult to understand, in fact the definition from NIH is simply the following:

Gene amplification is an increase in the number of copies of a gene sequence. Cancer cells sometimes produce multiple copies of genes in response to signals from other cells or their environment. The term also can refer to polymerase chain reaction (PCR), a laboratory technique that is used by scientists to amplify gene sequences in a test tube.

So my question is: if I have the complete DNA sequence of the organism's genome that I am studying (like this), that belongs to Archaea domain, what I should do to check for gene amplification is to see if the sequence of the gene (that I suspect that it is amplified) is repeated more times ?
I thank you in advance.

Could you clarify a few points to produce a focused question on the biology rather than the bioinformatics. (Of course a question on the bioinformatics is acceptable, but one at a time is how this site works. The two are separate questions.) When you say "how gene amplification is present in the genome" what do you mean? Do you mean how does it occur mechanistically, or what function can it serve? And it would be good to say whether you are interested in prokaryotes (as it appears) or eukaryotes, because the situations are rather different in the two cases.

@David ok I will correct it but with the  "how gene amplification is present in the genome" I mean how the genome DNA sequence is after gene amplification... so I am not interested in how it occurs mechanistically or what function it can serve (unless it is needed to understand what I want to know)... so in the sequence that I linked, should I simply find the sequence of a gene more times if it has been amplified ?

I appreciate the linguistic usage problems with the English word “how” for non-native speakers (French, German and Italian, rather than Spanish which I assume  is not too different). The usage you employed seems logical (= “how are amplified genes arranged/disposed in the genome?”) but is never employed. I would perhaps write something like “What is the typical arrangement of amplified genes in bacteria, especially those features that one might regard as indicative (or signatures) of amplification?”. It is not my particular area or I would answer.

Thank you very much @David for the suggestions

You ask "so in the sequence that I linked, should I simply find the sequence of a gene more times if it has been amplified ?"
Yes. There are many ways, both natural and artificial, to end up with more copies of a certain gene or genes without a proportional increase in copies of the other genes in the cell. Thus, there is no generally expected arrangement of the amplified genes.
One aspect to think about is the criteria for determining whether two given genes are copies of the same gene or are two distinct genes.

Thank you for your answer but I do not understand when you say One aspect to think about is the criteria for determining whether two given genes are copies of the same gene or are two distinct genes, because, if they have the same sequence, they should be certainly the same, right ?

I added a new post focused only on the second question, so if you want to have a look here https://biology.stackexchange.com/questions/101133/what-are-the-causes-of-gene-amplification-in-archaea you can understand better what I meant . Sorry for these jumpings :)

@Manuela Re: " if they have the same sequence, they should be certainly the same, right ?" Right, but many (most maybe) processes that produce gene amplification will result in mutations in one or more copies, either immediately or over time. For example, a duplicated segment of chromosome may initially have the same genes, but absent strong selection those will diverge over time. Maybe the copies still function the same, maybe they don't.

Thank you ! Things are much more complicated than I thought. Is there a criteria to enstablish if they are copies of the same gene, as you mentioned in the answer ? And, moreover, can you suggest some books or material that explains with more details what you said to me in the answer (so about the fact that we should see in the genome sequence different copies of the gene duplicated) ?

@Manuela My perspective is gene duplication in mammals over evolutionary time, so I would give an example of something like "Evolution of olfactory receptor genes in the human genome"  doi: 10.1073/pnas.1635157100 and the follow-on work. Your perspective seems to be much more short-term oriented (and not eukaryotic!), so something like "Bacterial gene amplification: implications for the evolution of antibiotic resistance" Nat Rev Microbiol. 2009 Aug;7(8):578-88. doi: 10.1038/nrmicro2174 might be more to your interest.


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Are all organelle lumens a reducing environment like the cytosol, or nonreducing like the extracellular space and the ER lumen?
I am interested to know if cysteine can form disulphide bridges in proteins within organelles. Typically cysteine will not form disulphide bonds in the reducing environment of the cytosol, but will in nonreducing environments such as the extracellular space.
From the Handbook of Biologically Active Peptides:

The ER is a vastly more common site than the cytosol (which is very rarely the site of disulfide bond formation) because the ER intralumenal environment is more oxidizing than that of the cytosol.

This is fairly intuitive since the ER is where the disulphide bonds will be formed before translocation to the extracellular space. But what about other organelles?
Does this apply to all lumenal spaces as implied in the below picture, or is the ER a special case as a result of its necessity to fold proteins destined for the extracellular space?

I can't speak generally, but I study a lysosomal protein that has many disulfide bonds so, at a minimum, the lysosome isn't reducing and I'd wager the entire endo-lysosomal pathway as well.


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Can 2 eggs come from ONLY 1 ovary?
I only have 1 ovary due to previous pregnancy and was told that, that pregnancy was a twin pregnancy. I know I can have identical twins but I wanna know the odds of fraternal twins.

It would seem to be obviously yes, since it's possible to have fraternal triplets: https://en.wikipedia.org/wiki/Multiple_birth#Triplets  I would guess that the odds for you having fraternal twins are about the same as a woman with two ovaries having triplets: about 1 in 1000.

The answer to this is yes, you can have fraternal twins. I don't know the likelihood of this happening, and I don't think there is any literature on it in the scientific community.
There is a phenomenon called superovulation, which is where more than one ova (egg) is released during a cycle. The classic model of one egg being released per cycle is still being investigated, and it is possible that it will be found to be true.
I had assumed (me not being an expert on fertility) that such a phenomenon was uncommon, but it seems that it is more the norm than not.
The most recent and most complete study I could find on this is an article on the development of follicles (pre-egg phase) in normal women from Canada. In the article they found that despite the current model of single waves of development and ova release, in most women there are at least two waves of follicular development per intraovulatory interval (IOI)1.

Nonrandom  changes  in  the  number  of  follicles >5mm and the diameter of follicles >6 mm (Non-Identity Method) were observed in all 50 women during the IOI, indicating a wave pattern of follicle development. Thirty-four of the 50 women (68%) exhibited two waves of follicle development; the  remaining  16  women  (32%)  exhibited  three  waves  of follicle development...  ...None  of  the  women  evaluated  exhibited  only  a single wave of follicle development during the IOI.

They didn't go so far as to say that each wave resulted in ovulation, despite several editorials claiming this was may be case in very reputable scientific journals (e.g. this British Medical Journal one).  In fact they say that only the final wave in each IOI is ovulatory (anovulation being without ovulation):

The final wave of the cycle was ovulatory and the preceding waves were anovulatory in all 50 women.

Of note is that for women who have undergone unilateral (one-sided) oophorectomy, the remaining ovary seems to compensate to some extent, producing regular menstrual cycles. I haven't been able to find out if there is an egg released from the one side each cycle or not, but I think most women don't necessarily alternate sides between cycles.

Baerwald et al., A new model for ovarian follicular development
during the human menstrual cycle. 2003. Fertility and Sterility 80(1).


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What are the causes of gene amplification in archaea?
I was studying this article about Genome Sequencing of a Genetically Tractable Pyrococcus furiosus Strain Reveals a Highly Dynamic Genome in order to try to extrapolate some features that could explain, at least in part, the survival strategy of Pyrococcus Furiosus exposed to gamma irradiation.
In this article, they compare the Pyrococcus Furiosus reference genome with the genome of a variant designated COM1, which is a genetically tractable strain.
In the Introduction they say:

The COM1 strain was obtained by targeted gene disruption of the pyrF locus (PF1114) using a plasmid designed for double-crossover recombination (37).

I would like to see if, after this targeted gene disruption, there is some amplification of some genes that I suspect they are involved in the servival strategy of these extremophiles to gamma irradiation, because then in the article they say that a number of genes involved in DNA repair could potentially be inactive in COM1 (as they fall into the minor protein-level differences category) but nevertheless it shows no significant changes in its ability to recover from exposure to UV or gamma irradiation.
For this reason, I would like to see if maybe some other genes involved in genetic material protection are amplified in response to this targeted gene disruption (in the sense that there are more copies of a gene being present in the genome). In fact, since I saw from NIH that:

cancer cells sometimes make multiple copies of genes in response to erroneous signals from other cells or because of signals they're getting from the environment.

Could this be valid also for my situation? In the sense that now I am not considering cancer cells but Pyrococcus Furiosus cells and as signal from the environment I consider the targeted gene disruption ?
Thus, my question leads to: what are the causes of gene amplification in archaea ? In few words, I need to understand this last question in order to see if it makes sense to try to detect gene amplification with the DNA sequencing data that I have. Anyway, I gave you all the background I am working with because I hope that, in this way, you can understand better what I want to know.
Thank you in advance.

Re: "what are the causes of gene amplification in archaea ? " I suspect trying to answer that would make for a good research career! :) I'm not familiar with archaea, but I think the NIH "signals" statement above only means that pre-existing signal-based gene amplification systems are induced in those cases.  However, perhaps Pyr. does have mechanisms to detect loss-of-function in a gene and respond as you suggest. Sounds like an interesting research project!

Thank you very much for your opinion :) ok I do not really understand the meaning of pre-existing signal-based gene amplification systems but knowing that it is not a completely nonsense reasoning is already something very important!  I think that there are few people studying archaea and I do not know why... They are so interesting, as you say .

The best science often comes from reasoning that gets close to "nonsense", but not quite. You've got an interesting idea to pursue -- good luck!

Just to clarify, are you interested in gene amplification (more copies of a gene being present in the genome), or increased gene expression (more copies of a gene being transcribed into mRNA and/or translated into protein)? In a pure culture system (clonal lineage), I think it makes more sense to look into transcriptional regulation before looking for gene amplification, though gene amplification can be favored at times (some forms of acquired antibiotic resistance in bacteria, for example).

@MikeyC I am interested in the first one: gene amplification (more copies of a gene being present in the genome). I have already studied gene expression... not really in this exact context. I made statistical data analysis of quantity of mRNA produced from some particular genes when P. Furiosus cell coltures were exposed to gamma irradiation. Now the idea is just to check if the genes, that in that study increased the mRNA production after the irradiation, are amplified if you do targeted gene disruption .

I really would have analysed this in the same context in which I studied gene expression... so for example to see the genome of the irradiated cell cultures and the not irradiated (control) and check if there is amplification of some particular genes in the irradiated one. The problem is that I do not have those kind of data. I can not find them. I do not even know if someone has done such experiment.

I see. Definitely an interesting topic. I know that viral insertion is one cause of non-essential gene duplication. Maybe looking for viral genetic signatures could provide some relevant insights. I can also imagine how viral excision or insertion might be triggered by a stress response to radiation exposure.

Ok, thank you @MikeyC . So maybe do you mean that some viral insertion mechanisms could be analogous to the targeted gene disruption...?


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Why is this Sea Turtle yawning?
I found this video online of a Green Sea Turtle lying down on a sea sponge and letting out a big yawn.  Here is a screenshot of the video, just incase the link doesn't work:

As far as I understand, Sea Turtles breathe air like we do, and effectively hold their breath underwater.  Also, as far as I understand, yawning is common to most vertebrates and it is believed to be triggered by the brain when there isn't enough oxygen in the blood.
If all of this is true, why is this Sea Turtle yawning underwater?  If it's not breathing the water like a fish would, surely the yawning wouldn't have the intended effect of putting more oxygen in its blood?  Is my understanding here correct?  And if it is, can someone please explain the benefits of a Sea Turtle yawning underwater?

The purpose of yawning is not clear, the "triggered by the brain when there isn't enough oxygen" explanation is just one and doesn't have much supporting evidence. It's also unclear to me whether this turtle is actually yawning or not, and I certainly don't think that can be assumed. Possibly someone else will be more familiar with sea turtle behavior, though.

@BryanKrause It does look like a yawn to me, but then again you are making me wonder if I am anthropomorphizing this turtle?

Bored with trivial questions on SE Biology?


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Why is there more female mortality in the middle of pregnancy?
NPR writes in Why Are More Baby Boys Born Than Girls? :

Scientists have found some unexpected clues that could help explain
why 51 percent of the babies born in the United States are male.
[...]
So that must mean the skewed sex ratio at birth happens during
pregnancy. Looking deeper, the researchers found that in the very
first week of pregnancy, more male embryos died, possibly as a result
of serious chromosomal abnormalities, which they also documented.
"When that settles out, it looks like there starts to be an excess of
female mortality," Orzack says. "And in the third trimester, as has
been known for a long time, there is a slight excess of male
mortality."

I suggest that you move this question to SE Medical Sciences, which seems to me both more appropriate and a place you are more likely to get an authoritative answer.

If you read the journal article referenced by NPR, the authors give some possible explanations for this phenomenon, including skewed X-inactivation and developmental retardation by the paternal X chromosome. See the second paragraph of the section titled "Overview".

It's worthwhile to note that the researchers were not able to assemble a uniform and random sample spanning all of pregnancy. Their information came from different sources during different stages of pregnancy and so is not directly comparable. Some of the sources are likely to be biased, e.g. amniocentesis is more likely to be performed if a problem is suspected.

The statement quoted in the question is hedging, a clear sign that its authors are uncertain of the reasons for the gender imbalance at birth.  As with anything in the scientific realm, there are usually more variables than can easily be considered or even filtered.
It used to be that about 106 boys were born for every 100 girls.  The numbers of boys are in the decline, a sign of loss in fertility.  This is most probably a result of environmental toxins, but may involve other lifestyle factors as well.
WebMD has a page HERE that details some of this (and the CDC concurs).  As its author points out, new mothers and young fathers tend to conceive more boys than girls--a clear sign that fertility is involved.
One theory, believed by many but still questioned by others, was promoted by Dr. Valerie Grant, Mr. David Rorvik, and Dr. Landrum Shettles (one article HERE).  It involves the physiology of sperm.  Biologically speaking, because the female X chromosome is very large, relative to the Y chromosome, "female" sperm are also claimed to be somewhat larger.  Their advantage is said to be endurance--being likely to survive in the womb longer than the "male" sperm.  But they are thought slower at swimming, particularly given certain environments.  Younger couples tend to be more active, and are more likely to have coitus near the time of ovulation, giving the advantage to the "male" sperm.  There are other factors involved, however, including the levels of acidity in the birth canal, and even how the mother's body protects and nourishes the sperm while waiting for ovulation.
As to why more females are thought to perish mid-pregnancy, the answer may be as boring as that it simply appears to be so because the mortality of boys at both the early stages and the late stages is higher.  Could it be, for example, that boys are more prone to getting strangled by the umbilical cord?  There are some distinct gender differences with respect to the development of the umbilical cord and placenta, including increased risks for boys in the latter period of gestation, as noted HERE.
Even though more boys are born, boys have a lower survival rate.  They can be more accident prone, more likely to succumb to sickness, or to have more impairments from birth defects.  By the early teen years, the male-to-female ratio has usually corrected itself, and by the twenties there are more females in the population.  One chart providing a quick overview of this information for US populations can be seen HERE.

I'd suggest deleting everything after the first paragraph. The 1st paragraph is good, but the rest is not really scientific.  What exactly is meant by a "loss in fertility" and why would that affect the sex ratio at birth? There's no verified link to "environmental toxins" whatever they might be, and so forth.

@Armand If you had followed the links I included, some of your questions might have been answered.  Environmental toxins include elemental toxins like lead, cadmium, mercury, arsenic, bismuth, and antimony as well as biotoxins from other sources.  They come into our environment in many ways, and there are studies showing how they affect fertility.  Toxicologists can quantify the levels of toxins in the body through laboratory tests like a provocative urine test, a hair-mineral analysis, and a newer technology that uses a laser to scan the skin and spectrum analyze it with computer support.

my point is that such "explanations" are so nonspecific as to not be scientific. There is no credible science that I'm aware of indicating that typical exposure levels of any "environmental toxin" affects the sex ratio at birth in the US. Again, the answer says "fertility is involved" - what does that mean? The sperm X or Y content affecting fertilization rate has no credible evidence. The umbilical cord and last paragraph issues seem out of the scope of the question, and the increased mortality rate in the 1st half of pregnancy is real, not an illusion, according to the data shown.


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Do crabs actually "rip the legs off of other crabs" if they try to escape a bucket?
I read this strange remark:

Picture a bucket full of crabs with no lid. The moment a crab tries to get out, the other crabs will literally rip its legs off.

It was said as something that is common knowledge. I had no idea about this. What are they talking about? Huh? Why would crabs behave like that? Is this just nonsense?

No, that's an exaggeration of the "crab bucket" phenomenon.  They just pull the other crabs down: https://en.wikipedia.org/wiki/Crab_mentality  It's also a metaphor for a certain type of human behaviour.  See for instance Terry Pratchett (in Unseen Academicals): https://wiki.lspace.org/mediawiki/Crab_Bucket

@jamesqf Well, I watched numerous videos on YouTube but couldn't find a single one where this actually happened. Very frustrating.

I've no personal experience of live crabs, so I can't say whether it actually happens.  But as a metaphor, it's perfectly understandable.

As lobsters are crustaceans that I am more familiar with, I will say that lobsters are cannibalistic at times, so in addition to aggression, crabs may also exhibit the same predation behaviors when in close contact. I am not sure if that explains this behavior, but it is possible. Both crabs and lobsters do randomly bite each other when trapped together though.

Thank you for your enthusiasm, but please review the criteria for [answer] — in particular we want answers that provide detailed explanations with references to reliable sources (preferably primary literature). Speculation that isn't backed by such sources is indistinguishable from opinion, which is expressly off-topic on this site.


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Are there regularly spaced connections between caudate nucleus and putamen?
There are a number of images on the Web, most beautifully Visible Body's artwork, which show the caudate nucleus and putamen linked by regularly spaced connections across the barrier of the internal capsule.  Since vertebrate forebrains are treated as having only a few segments (see Michael Carstens, 2017), which seem tough to relate to the C-shape of deep cerebral structures, there is something about such figures that piques curiosity.  In a paper like Heide Hörtnagl et al., 2020 the situation seems less clear to me, but they weren't really focusing on this issue.  Is there a regular/regulated spacing between the connecting bundles, and does it correlate with any other observable anatomical regularity?

"vertebrate forebrains are treated as having only a few segments" - I think you may be overextrapolating from early developmental models (especially those making cross-species/evolutionary comparisons). In any event, that structure wouldn't have any relationship to the pattern I think you're asking about.

These seem to be referred to in literature as "transcapsular" or "caudolenticular" (gray) bridges. My initial thought was that the picture you shared is just some weird artistic rendering but upon seeing histological slides these are quite prominent and familiar to me as part of the basic "look" of these structures in histology. You can see them in any coronal slice through this area, this atlas, "Cross-Sectional Atlas of the Human Head" by JS Park has them labeled though often they are not labeled.

(left image from JS Park; annotations on the right image are mine. There are many more "bridges" than I've drawn but I highlighted the most prominent to compare on the left side. Probably those deeper in the section relative to the camera are showing up a bit faint or blurred. This is also just one histological section, the bridges are found throughout the length of the area where caudate and putamen are located, at least a couple cm)
There may be other terminology, as well, but I'm having trouble finding it.
I see no reason or foundation on which to connect these to any sort of segmentation of the developing vertebrate forebrain. Since resources are sparse it seems difficult to find a reference, but my thought is that this is just an emergent pattern you get as different fiber pathways cross one another, like how your fingers look when you fold your hands together.
Some other references:
Mendoza, J. E., & Foundas, A. L. (2008). The Basal Ganglia. In Clinical Neuroanatomy: A Neurobehavioral Approach (pp. 153-193). Springer, New York, NY.
Nieuwenhuys, R., Voogd, J., & Van Huijzen, C. (2008). Brain slices. The Human Central Nervous System, 137-173.

Thank you for a great answer!  Though perhaps I should have asked a better question...  I don't have ready access to these resources - can you tell from the sections you see whether the number and positioning of the bridges is consistent between individuals, as you'd expect from a regulated structure, or does the number and position vary more randomly as you'd expect from something emergent but unregulated?  Thanks again.

@MikeSerfas In the limited experience I have with anatomy of individual human brains (albeit focused on cortex) I have to say I'm amazed by just how much variability there is. I can't see these structures as having consistent structure but don't have a source to prove that either, only a strong suspicion that if they did have a regulated structure it would be remarkable and highly discussed. I would say though that the artist picture you show vastly underrepresents the number which may be why they appear so orderly.

@MikeSerfas I added a picture of one of the slides I referenced. I think you'll appreciate that they are quite diffuse and scattered, not nearly as orderly as that cartoon suggests, nor are they found in a plane like it appears there.

Oh!  I have seen those in images, but based on the artist's drawing I was expecting larger, rarer connections.  Now I see what you mean.

@MikeSerfas Yeah that was exactly my thought too - went from something I hadn't heard of to seeing a picture and realizing I recognized that from any coronal section of the the caudate I'm quite familiar with the look. I don't know for sure but I'm guessing it's that those fibers are unmyelinated/less myelinated than the rest of the fibers.

@Bryan Krause
Are these caudolenticular bridges not gray matter structures? So, they're not myelinated.

@Akila Not myelinated or at least less myelinated/lower total fat content than the white matter they interleave with.


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How does protein knockdown allow a phenotype to be observed?
I am trying to understand how AID2 technology works.
I came across an article that says "Protein knockdown using the auxin-inducible degron (AID) technology is useful to study protein function in living cells because it induces rapid depletion, which makes it possible to observe an immediate phenotype".
Can someone please explain in simple terms how one could make inferences about the phenotype based on protein depletion via AID2.

Here's a starting point for some research: "Auxin-Inducible Degron System for Depletion of Proteins in Saccharomyces cerevisiae"  PMID: 31503416 PMCID: PMC6741457 DOI: 10.1002/cpmb.104 One way to study protein function is to see what phenotype you get without that protein. That can be difficult if the protein in question is essential for the cell to grow or survive. This system allows you to grow cells normally, then at a time of your choosing add auxin and trigger the cell's own protein removal system to get rid of the protein of interest.

Phenotype is defined as the observable physical properties of an organism (organism here includes things like cultured cell(s)).
If you knock out the protein of interest you may observe a change in the function, behaviour etc. of the object you are studying via the methods you are using to look at that system. For example, in terms of proteins you might be looking to see how removing a certain protein affects production of a metabolite to investigate which pathways are being for production of that metabolite.

Yes, the "phenotype" is just what you observe, no "inference" involved.


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Could you identify the species of this animal attaching to a fish in the Aegean Sea?
While wading within a few meters from the shoreline today, I observed and was observed by a number of curious fish. One of them had a blackish attachment, which was reminiscent of a small aquarium catfish, probably because of its size and because its colour was not homogeneously black and appeared to have dense brownish spots. But it had no apparent thin parts like fins or tail and was quite plump overall. For this reason, I do not believe that it could be remora or lamprey, but could it be a marine leech? It was probably less than an inch long and one third of that wide. Could you identify what species could it be considering that it was in Toronean Gulf to the south of Kallithea village? I attach a collage of somewhat blurred photos captured through the surface of the water.

Update: I've seen three more examples today in the same spot. No pictures this time. The host fish species was the same. The parasite looked the same. The rear part of it is narrower and thinner than the rest of its body and the rear edge is uneven in comparison to the rest of its contour. In three cases out of four, it was attached to the same place at the right side of the fish. In one case, it was attached to the same place but on the left side of the fish.

Given the rough shape I lean towards an ectoparasitic isopod. I have no idea which one of the roughly 2700 species it might be, but I would guess from a quick search that it probably belongs to the families Cymothoidae or Gnathiidae as these seem to be the more common and well described (at least they have more complete Wikipedia entries, wheras the other families don't).

Looking at the parasitic isopods which were reported in Aegean Sea (per Kirkim F, Kocataş A, Katağan T, Sezgin M (2008)), I lean to Anilocra physodes because of its overall shape, dark colour without longwise stripes, attachment to the rear side of the fish ("Caudal peduncul" per Öktener A, Alaş A, Türker D (2018)). The fish looks like Lithognathus mormyrus, which is a common host for it.

Gnathiidae ("fish gnats") would be 1–3 mm (see Williams E, Bunkley-Williams L (2019)).


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What is this bird similar to a seagull but with a forked swift-like tail?
I observed these birds in Saint Petersburg, Russia this June, mainly in the evening when the sun was setting. They are similar to seagulls and often fly (presumably hunting) at the same location as the seagulls.
Easily noticeable differences from seagulls are:

Forked tail, like swifts have
Narrower wings, which seem longer
Higher-pitched short tweets, which never turn into "laughter". A sample (with car noise in background) can be heard here.

Here are some photos I could took. They are blurred and chromatically aberrated because it's hard to focus with a telephoto lens and such fast motion towards and away from me. I exposure-compensated them to make the colors visible, so the photos are a bit noisy. I hope they are useful for identification anyway.

It would be helpful if you could [edit] in data about size (e.g. by comparing them to the size of birds at the same hight that you can identify). At very least that would allow you to rule in or out the little tern, which is about 2/3 the size of the other reasonable candidates.

This is one of the numerous species of tern. They are common birds of the seashore and rivers world-wide. The characteristic forked tail and the black cap on the head is what gives it away.
It is hard to say for sure given the chromatic aberration altering the colours, but this is likely to be either from the genus Sterna (particularly the Arctic tern or the common tern), or possibly the little tern (Sternula hirundo).

Looks like an arctic tern. The translucent wing feathers are easier to see in this out-of-the-cam photo without exposure compensation.

@Ruslan Seems most likely given you were in northern Russia, and the others seem to be in more temperate ranges.

"Gull + a forked tail" = "tern". It's hard to tell from your pictures, but: Common tern: red bill with black tip. Arctic tern: red bill with no black tip.  Roseate tern (rare): black bill. Sandwich tern: black bill with yellow tip. Little tern (which is really small compared with the others): yellow bill with black tip: Most juvenile birds (of all species) have black or gray bills.


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Reference for standardized histology rating criteria of microscope images
I am looking for a reference to a standard protocol for assessing the visual quality and diagnostic potential of a tissue sample that has been processed and stained for viewing under light microscope. What kinds of things are important to look for? For example the overall morphology, cellular outline, integrity of structures, nuclear details, lymphocytes and erythrocytes, etc...
I am looking for a paper that explains what things to look for in a well processed sample. Thank you

I think this question would be a better fit in Biology.SE because histology is much broader than medicine.

Consider the 1951 USAF resolution test chart (inline below). It has arrays of progressively smaller spatial frequency features. Just find the smallest feature that can be resolved and use this as a metric to compare to other brightfield microscopes in your facility.

Such charts are available from Thorlabs, Newport, and Edmund optics. Edmund also has a calculator to use with it which provides the resolution in line pairs per mm.

I have edited your answer to direct include the image and a link to wikipedia for the item, and the additional info you provided by comment. This is the kind of thing that I meant. Also note that while you have addressed issues of resolution, you have not really addressed the more relevant (IMO) biological side of the question.


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I already heard filtering the air to get rid of the Virus is not that effective but what about UV radiation?
Why they don't add a chamber to their air conditions and give the air treatment with UV radiation? I'm not a science guy. Please what might be the reason? Wouldn't it be also cheaper because the air conditions don't need to pressure the air through tiny filters?

I’m voting to close this question because this is not a forum or tutorial service; it is a Q&A site where specific questions relating to biology get specific answers. Please take the [tour] and carefully read through the [help] to learn more about the site, including what is on-topic and what is not, and how to [ask] a good question. See also the section on "homework".

UV is highly effective against many viruses and bacteria . It needs to be germidicidal UV, often known as UV-C in the short UV wavelengths, so there is no doubt that this could work (indeed there are a number of people working on it at the moment, and even some devices available). This sort of technology has been in place in healthcare settings for disinfecting patient rooms for immunocompromised cancer patients for example.
The difficulty with this is providing enough UV to the air-flow so that it deactivates the virus during the short time the virus is passing the UV bulbs. There's a good source of information at the Illumination Engineering Society about UV and germicidal activity here (PDF), which specifically discusses SARS-CoV-2 and the mechanics of how to do it. It discusses in terms of air-changes-per-hour (ACH) and dose for a number of situations:

... simpler evidence-based dose that has been developed over many
years for TB control, typically specified as about 17 mW of 254-nm lamp-emission radiant power per cubic meter (m3) of space to disinfect air.

17 milliWatts of power might not seem like much given that your room lights probably use between 5-200 Watts per bulb, but they are putting out a broad range of light (mostly as heat) from about 400 nm to 800 nm, but the UV bulb output that is being measured here is the specific germicidal wavelength of 254 nm, overall the lamps put out much more light than that and are enormously energy hungry.
It also says that disinfecting the volume above people in a room is more effective than irradiating incoming (or presumably outgoing) air because of the risk of person-person transmission with people in the same room:

Upper-room GUV (see Figure 3-1) disinfects large volumes of room air (above occupants’ heads) at once,
resulting in high "equivalent" air changes per hour (ACH) in terms of air disinfection only...
...When UV is used in ducts, although it ensures that recirculated air does not have viable pathogens, it
unfortunately does relatively little to prevent person-to-person transmission in a room where both an
infectious source and other susceptible persons share the same air. For effective interruption of
transmission, air disinfection has to occur in the same room where transmission is occurring


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Methods for getting clean C. difficile spore preps without spores clumping together
I'm a microbiologist recently who recently started working on an ongoing C. difficile project for the first time. One of the things I need to do is verify the lower detection limit of my qPCR detection assay (C. diff Toxin B gene) using C. difficile spores spiked into negative clinical specimens. I also need to obtain accurate viable spore CFU counts in order to generate the appropriate spore titers for animal experiments. I'm trying to get a clean spore prep free of vegetative cells (for the spore detection assay), and minimize the number of spores clumping together to obtain more accurate CFU counts (via plating heat-treated spore dilutions on BHIS+taurocholate germination plates).
I'm working with the C. difficile strain 630 (ATCC BAA-1382), a lab strain that's notoriously not the best sporulator (we use 630 because it's toxigenic and infectious, but not highly virulent. This helps minimize animal suffering in simple infection models where we don't need to look at toxicity).
There are a few commonly cited C. difficile spore prep methods that I've been working from, and in some cases trying to mix and match possible solutions from different protocols, but none of them were use this particular strain in the methods papers. Our lab SOP uses Clospore medium, and that seems to give me the best sporulation rates compared to the solid 70/30 medium used in the STAR protocols paper. But when I follow the rest of the Clospore protocol for spore collection and purification I end up with highly concentrated vegetative cells and am getting lower viable spore titers in the "concentrated" spore prep than there are in the unconcentrated cultures (by multiple orders of magnitude). The spores I do recover are mostly clumped together into large clusters when viewed under phase contrast microscopy (possibly explaining the lower spore CFU titers).
If anyone has experience with getting really clean C. difficile spore preps with minimal spore clumping (especially in 630), I would love any tips or advice.
Potential solutions that I've tried without success:

Saving the pellet wash supernatants and re-pelleting with a longer centrifugation: This does recover some spores with almost no vegetative cells, but they are very low titers and still have clumping issues.
Using room temperature water instead of cold water for washing the spore pellet, (it's been reported that cold-water wash induces more clumping): Mine still clump with room temperature water, and there's no difference in terms of vegetative cells.
Separation of spores from vegetative cells using a 50% sucrose gradient centrifugation: This does separate out a lot of cellular debris, but the intact vegetative cells pellet out with the spores, and still has spore clumping issues. Plus I'm losing a lot of spores when I do this (possibly due to extra wash steps needed).
Washing with 0.05% TWEEN/Polysorbate (nonionic surfactant): This actually helps to free up spores sticking to the culture tubes, but they still clump together when washed to remove the TWEEN (polysorbate has toxic effects in mice, so washing it away in not optional).

I’ve had similar poor luck getting sufficient spore concentrations following this protocol and using ATCC strain 9689. Lots of vegetative cells, few spores. One recommendation from that protocol is to use PBS + 1% (w/v) BSA to resuspend spores after washing to prevent clumping, per Edwards et al.

@acvill Thanks! I'll will try that for sure.


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Does DNA have 5 nitrogenous bases?
Does DNA have 5 nitrogenous bases? I believe they are 5 because Uracil is not the same thing as Guanine, because, first of all, uracil "replaces" thymine, not guanine. And second, uracil performs a range of functions that go far beyond "replacing" thymines at the time of transcription and translation. Am I right?

No. Uracil is present in RNA which is not the same as DNA.

The Biology.SE community has agreed that questions that show little or no prior research effort are off-topic on this site as are "homework" questions unless you have shown your attempt at an answer. Please [edit] your question and tell us where you've looked for answers, what you do know about the topic, and where exactly you still have questions. Under researched questions may be subject to down-voting and closure. In particular, I have a found KhanAcademy to be a very helpful introduction to unfamiliar subjects.

Thymine is 5-methyl-uracil, and one way of remembering this is that THYMinE and METHYl are (almost, but not quite) anagrams.

Uracil in DNA arises from the deamination of cytosine , and as such replaces C and not thymine. Thus a C-G base-pair is converted (by deamination) to a mutagenic U-G base-pair (and there are sophisticated repair mechanisms to fix this).  See Uracil in DNA – occurrence, consequences and repair, for example

It's a bit more complicated than that.
This wiki entry on nucleotides should help clarify things: Nucleotide
Diagram of chemical structure of bases, nucleosides, nucleotides, etc
I'm not sure what you mean by "...a range of functions..."

From How do I write a good answer? in the [help], Answer well-asked questions. Not all questions can or should be answered here. Please save your efforts for questions appropriate to this site. I also encourage you to delete your answers to  such questions (e.g. homework questions like this) as giving them answers encourages more undesirable posts. This is also destructive to the learning process by rewarding lack of thought and effort on the part of students. Thanks.

For "It's a bit more complicated than that", you could reference Uracil in DNA – occurrence, consequences and repair. (Uracil in DNA arises from the deamination of cytosine)

@user338907 I was attempting to provide a starting point for the questioner's research rather than a direct answer, as suggested by (at)tyersome (I may not have succeeded in doing so).


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Understanding Contig NG50
My goal is to do a de novo assembly of a jellyfish. At the moment I'm busy checking papers that did de novo assemblies but I also got some jellyfish papers
Then I stumbled accross this:

Contig NG50 is defined as the length at which half of the predicted genome size is contained in contigs longer than this length.

I can’t understand this and googling returned papers of similar complexity e.g.:

NG50 refers to the length of the smallest contig added to cover 50% of all nt estimated in the
genome; (25)...

Can someone provide a simpler explanation?

Please do not direct users of this site elsewhere for the meaning of specialist abbreviations central to your question. Questions here should be self-contained, and if you request clear answers, I suggest you do as you would be done by.

The Wiki entry "N50, L50, and related statistics" would be a good research resource. https://en.wikipedia.org/wiki/N50,_L50,_and_related_statistics

This stack is for professional/student biologists, not 5 year olds, so I'm unsure why you seem so insistent on on dumbing down the title.

@user438383 Because my professors always told me only truly understand something if you can explain it to your 5 year old brother :) And also because I thought this is a positive and supportive place where people are encouraged to ask questions that other people are encouraged to answer (collecting votes I assume). Have you ever read a paper that was not in your native language? It can get very tough, but I hope that I understand a little bit more every day, with the kind help of smart and motivated people like you and other members of this community, but maybe I'm just a naive dreamer :)

A title SE should indicate the biological content of a question. It should be totally impersonal. Casting it as a challenge to users is against the ethos of this site, and your particular example is made worse for the reasons in the comment made by   @user438383. We welcome new users to this site, but we expect them to follow the norms set out in the Help. Please do so.

I have edited your question to provide a partial model of good practice. (The first sentence is too chatty.) If your aim is to communicate the results of your (genome?) assembly you would do well to study it.

If you arrange your contigs from longest to shortest, what is the shortest contig in the smallest subset of contigs that covers at least half the expected genome length?
An example with arbitrary units...
You have a set of contigs of lengths 30, 20, 20, 10, 5, 5, 1, 1, 1, and many contigs < 1.  Your expected genome length is 150. The smallest subset of contigs that covers at least half the genome length is [30, 20, 20, 10]. The smallest contig in that set has length 10, so NG50 = 10.
Note that the sum of the lengths of the contig subset is 80, not 75. But if you take away the smallest contig in that subset (10), then the sum is 70, which is less than half the genome length. So the sum of the lengths of the contig subset need not be exactly half the expected genome size, but at least half.


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Relative abundance of transcription factors and protein kinases
Are transcription factors and protein kinases only expressed at low levels in eukaryotes?
As regulatory proteins, I would expect their abundance to be lower than most other proteins, but I cannot find any published research to support that.

@l0110 Replacing makes the question "Are transcription factors..."

You are grouping genes by overall function "important cellular regulators" but asking a question about a putative shared characteristic "expressed at low levels" for genes which act via very different mechanisms. I would think looking into their mechanisms would help answer your question.

Summary

The abundance of protein kinases and transcription factors vary among and between these broad classes of protein.
General Considerations
Let us first think what might be expected for the abundance of transcription factors and protein kinases.
Protein kinases: These would be expected to be less abundant than their substrate proteins as they are enzymes catalysing a reaction. However some protein kinases are part of amplification cascades (e.g. MAP kinases) so the abundance of individual member would be expected to differ.
Transcription factors: Transcription factor is a portmanteau term that includes both proteins that act generally on promoters (e.g. the TATA-binding proteins TFIIA, TFIIB etc.) and ones that are specific for the promoters of particular genes or classes of genes (e.g. heat shock factor). The abundance of transcription factors might be expected to be inversely proportional to their specificity.

Correlation between Protein Kinases and Transcription Factors: I can see no reason to group these two classes together as the regulation of metabolic activity and gene expression are very different processes. All I would expect is that they would to less abundant than structural proteins such as actin or ribosomal proteins (although differing in abundance among themselves).
One Possible Approach

To the man with a hammer, everything looks like a nail. My particular hammer may not be the best tool for this particular nail, and there are certainly other tools available which other answers may mention, but it’s something you can do at home (without even adult supervision). The hammer is called FlyAtlas 2 and can be accessed at http://flyatlas.gla.ac.uk/FlyAtlas2/index.html. It allows one to examine the extent of expression of different genes in the tissues of the fruit fly, Drosophila melanogaster.
Some Words of Caution

Before looking at experimental data, one should be aware that the abundance of enzymes will vary depending on the particular tissue examined and the physiological conditions. One should also be aware that the data are for abundance of mRNA transcripts, which may not always be proportional to the amount of the encoded protein. Further, although the resource has been published in a peer-reviewed journal, as is normal in science, the referees have not repeated the experiments.
How to Proceed

Connect to http://flyatlas.gla.ac.uk/FlyAtlas2/index.html
There are two ways of proceeding here. One can go to the Category page, type in a term in either as a Category or Free Search, and select from the menu, as shown.
Alternatively go to the Gene page, click (or tap) on Gene Name and choose from the autocomplete. This gives fewer results than 2, but they tend to be grouped in a way that makes it easier to make repeated searches.
Click Search and tick Whole Body in the results (which may be downloaded in a spreadsheet-friendly format). An example is shown.

Results

A quick application of this approach shows that:

Protein kinases are expressed in whole flies in a ten-fold range, from just detectable (e.g. Cyclin-dependent kinase 2) to 20–40 FPKMs (protein kinase A, calmodulin-dependent protein kinase).

Some protein kinases are highly expressed, but only in specific tissues (e.g. Erk7 in testis).

Transcription factors are expressed in whole flies in a range from just detectable (e.g. Transcription factor AP-2) to about 10 FPKM (TF IIB, heat shock factor)

Some Transcription factors are tissue-specific (e.g. TF B4 in ovary and testis).

Comparative values of cellular enzymes are glycogen phosphorylase (c. 100), phosphofructokinase (c. 50), enolase (c. 200).

Thanks David. I usually use R or Python for these analysis but it is nice to know there is a website to do it. My own analyses showed these two groups of genes are expressed at low levels for mRNA and protein when comparing to other groups of genes in plants. Since I want to know if there is a reference there (that was the original question), I up-voted your answer but not accept it. But thanks again for your time and effort!

@l0110 The website is, like others of its type, a front end to a relational database, providing the general user with the ability to make a restricted range of queries. It also provides APIs for the programmer to make queries in whatever language he prefers (see online Docs). Although perhaps not of interest to you as a plant scientist, queries using the recent categorization of fly genes into classes on FlyBase might be a better way to approach the question. I imagine there are similar resources for Arabidopsis. Btw. I have deleted my comments on your question now.


text:
Clarifying Resting membrane potential
I need some help understanding how the resting membrane potential is maintained in neurons. I understand that there are more K+ leak channels than Na+ leak channels, meaning that more K+ ions leave the cell membrane than Na+ ions enter.
However, where I am confused is with Na+/K+ ATPase. 3 Na+ are pumped out and 2 K+ are pumped in. From what I understand, this pumps out Na+ as fast as it leaks in. But if K+ leak out faster and less are brought in, then won't there be eventually no K+ left in the cell?

Better to not worry about the pump ratios at all. They don't matter appreciably. Learn and use the Goldman/GHK equation.

Think of it this way, the rate at which K+ and Na+ try to leak to the side with the least concentration  of that ion  is counteracted by NA+/K+  pump. So while K+ is leaking out and Na+ is leaking in, this process is balanced by the pump (pumping K+ in and Na+ out). This is process will come to a balanced equilibrium, where ions will still be leaking, and still being pumped, but the net sum of these two opposing processes is zero change in concentration.


text:
Eye vision for distant large objects versus small close objects
We often hear that looking at distant objects after staring at screens is good for the eyes to prevent strain.
But from the perspective of eye muscles, what is the difference between small font that is up close on a screen and a poster with large words on it say 30 meters away?
Since if something grows in size as it travels away from you - from your perspective theres no way you could perceive it as moving away from you.
Doesn't the eye still have to focus in the same way, so why then why would looking at objects 30 meters away act as a way to counter strain from looking at smaller font closer up if they are two sides of the same coin?

Welcome to Biology.SE. Please clarify your question. It seems to be about focal length, which is really a physics question. Perhaps you are trying to ask about how the eye changes focal length — if so, please do some research on your own and then ask specific questions about what is still unclear to you. Also, it is unclear what perception of movement (paragraph 3) have to do with the rest of your question. In general, questions on this site must: demonstrate prior research, be clear, be specific, and be about biology as defined in the [tour]. ...

... You will have a better experience on this site if you familiarize yourself with the expectations by consulting the [help] starting with [ask], what is on-topic, and what is not. Thanks.

I don't see how this question does not fit here even after reading those links...its not about focal length in terms of physics its about how the eye perceives distance in order to adjust its focus. How can the eye tell the difference in order to make the statement that looking at long distances relieves eye strain from looking at short distances from the example i gave.

I did but I am not finding any solid answers other than physics equations but thats not really answering how the eyes/brains interpret it. @Laura_Jonees

I suggest you do some background research on optic first because your premise is way off.

@John no idea what you're on about with my "premise" being way off. I have a degree in physics and know full well about lens equations and optics. But that has absolutely nothing to do with what im asking. Since i am asking about biology and neurology - not physics. But thanks for your contribution - which was basically nothing. IF you know something then provide your knowledge or your comments are unwanted here. Thanks.

the size of an object has no effect on its focal length, so review your basic knowledge of optics. A big object at 30m and a small object at 30m has the exact same focal length and needs the exact same lens to be in focus. this may help https://www.stemmer-imaging.com/en/knowledge-base/optics-technical-overview/

Did you read my question, i am not talking about two objects at the same distance... so a big and small object both 30m away is not what i'm asking. I'm asking about a small object close to the person, and a large object far away. I don't need your physics answer this is a biology question.


text:
Cyanobacteria and the ratio of Iron to Phosphorous
In listening to a recorded lecture that touched on the Great Oxidation Event, a casual remark was made to the effect that the shifting of the Earth's constituents to more availability of phosphorous relative to iron was a key factor in the ability of cyanobacteria to produce oxygen.
[Although it was primarily a geology lecture, it did not mention what caused that shift.]
How would this change (if one could go so far) prime the pump for the GOE?
Thanks

https://www.frontiersin.org/articles/10.3389/fmicb.2020.541558/full

@MaximilianPress Thanks. Hardly "In"!


text:
How could a water boatman bug get in a place with no water?
I built a pond in my backyard. Before I even topped it off, I started finding water boatman (Corixidae) swimming in there.
Note that the nearest body of water from my home is 30 miles away, and its about 1/2 acre in size. There are dry river beds, dry canals, and even a dry lake nearby, but I've never seen water there in 5 years being here.
So how is it that water boatman can show up so suddenly in my pond?

In animals some species migrate, some hibernate and some aestivate to
overcome the unfavorable conditions such as desiccation.
Some aquatic organisms enters into diapause during unfavorable conditions.

Source: toppr, also see Gemmule (in sponges)


text:
What species left these underwater eggs?
While inspecting some plants planted about 1.5 ft underwater in my pond, I noticed what appeared to be eggs on many of the plants.
Note that the eggs are the white, oblong, opaque things. The clear perfect spheres in the picture are just water bubbles.
For scale, the branch is probably 1/8" thick. These are really tiny, maybe 1 mm across.
The location is the American southwest.
The eggs had to be at least 1 ft underwater, which rules out most kinds of flying insects, I think. What kind of animal lays an egg like that?


text:
Why are heterologous expression systems such as mammalian cell lines commonly used for studying interactions between neuronal proteins?
I am reading papers about heterologous expression systems and I have seen that they are used a lot in neuroscience to studying interactions between proteins that are normally expressed in the brain.
I have seen that in research papers, CHO cells are used as a model cell line to study the interactions between neuronal proteins. This is done through overexpressing a protein of interest that is natively expressed in neurons in CHO cells and investigating interactions of that protein with another protein (that is endogenously expressed in CHO cells but also found in neurons).
However, I was wondering, what are the advantages of using CHO cells instead to study interactions between neuronal proteins instead of a cell culture of neurons? Wouldn't it be more accurate to study interactions between neuronal proteins using cultured neurons instead? Any insights are appreciated.

Should depend on the interaction being studied. If the interaction happens within a single cell a cell line should suffice.

@ceno980 I do not know why your question is being downvoted. I feel this is a genuine question researchers come across when they getting into new arena.

One of the advantages you already stated that depending on protein of interest if it's already expressed in CHO cell line, researchers use that.
Apart from that, this question is more of a general type. I will try my best to logically put down the things.
You are right, it's more appropriate to study proteins using cultured neurons. But, there are many practical factors that may compel us to avoid cultured neurons.

Ease of Culturing cells
Its way easier to culture cell lines such as CHO/HEK293/HeLa compared to neurons. Neurons are very sensitive and culturing them needs serious expertise and still, things can get wrong. There are many labs across the world that have dedicated technicians to avoid individual differences between cultures which can topple up experimental results. In contrast, culturing common cell lines doesn't need that level of expertise (relative comparison). Furthermore, for neuronal culture, several serious ethical guidelines need to be followed and require a good amount of funding to set up the facility. In addition, to have neurons develop processes and ready for study generally 10-12 days are required which prolongs the length of study. With common cell lines, you can perform an unlimited number of experiments and get results in a relatively short amount of time.

Ease of expression
Sometimes, the endogenous level of expression is not sufficient enough to visualize the interactions. More importantly, you do not have a tool to visualize these endogenously expressed proteins in cultured neurons. At that point Transfection and overexpression comes in handy. Transfection and overexpression in cultured neurons are way more difficult compared to common cell lines such as CHO, HEK, HeLa, etc. The efficiency is very low. It needs serious optimization and still, everything may go in vain. If endogenous expression levels are sufficient enough, knock-in expression is an option. Still, it requires several months to have such knocked-in mice and the success of knock-in is very low with larger proteins such as fluorescent proteins. Indeed new techniques are getting developed such as viral methods etc. Still, they are not well established across the globe.
In the case of common cell lines, it's relatively easy to transfect/optimize the conditions to obtain abundant protein of interest for the study.

Thus weighing time, money, expertise, and whether or not cultured neurons absolutely necessary to draw meaningful conclusions generally determine which cells to choose.

Thank you for your reply. Regarding your statement: "More importantly, you do not have a tool to visualize these endogenously expressed proteins in cultured neurons." I was wondering, is it not possible permeabilise the cell membranes of the neurons and then use indirect immunofluorescence to detect proteins that are endogenously expressed (even if they are expressed at a very low level)?

(1/2) Good question.There are some reasons for this.Let me clarify one thing first. Endogenous doesn’t necessarily refer to proteins present inside the cell rather it means ‘natively expressed’.Not always you want to permealize and visualize intracellular proteins. For example 70% of the FDA approved drugs target membrane proteins. Coming back to your exact question,currently we don’t have antibodies for many of the proteins of interest.Cross reactivity,low affinity etc are the common issues with available antibodies.

@ceno980 (2/2)(continued) Therefore you see studies genetically attaching fluorescent proteins/epitope tags to the protein of interest to study/visualize them.Still they are not perfect. That’s a different topic for discussion.

Thank you, for your reply it really helped


text:
Tips for longer fragment size and higher purity of insect DNA
Aim
In a pilot experiment I tested three different genomic DNA extraction methods on the non-model organism Pieris mannii (southern small white; Insecta, Lepidoptera, Pieridae). The goal was to generate DNA fragments of ≥30 kbp average length for library preparation for PacBio SMRT long read sequencing  (HiFi)).
Methods
The three methods/kits used were: Qiagen blood & tissue kit (column assay), Beckman & Coulter DNadvance  (magnetic beads kit) and standard Phenol-Chloroform extraction.
Laboratory raised organisms in the prepupal stage (termination of larval growth phase, after last excretion of intestinal content, no chitinous exoskeleton) were harvested and frozen at -20°C. For the DNA extractions the head and the first two leg pairs (ca. 20mg) were cut and processed according to the kit/standard protocols with some modifications for high molecular weight DNA:

avoidance of freeze-thaw cycles
pestles were used for grinding the tissue (no liquid nitrogen)
inverting instead of vortexing
cell lysis and proteinase K digestion at 56°C and 300rpm overnight
planned: use of wide-bore pipette tips (impossible because of very long delivery waiting time)

Results summary

Method
Avg. DNA fragment length (bp) range
A260/280 range**
A260/230 range**
Nanodrop avg. conc. (ng/µl)
Qubit avg. conc. (ng/µl)

Qiagen
1038 - 18366
2.07 - 2.12
1.77 - 2.15
1151.8
104.9

Magnetic  beads
7350 - 12586
1.98 - 2.11
1.60 - 1.76
1048.7
155.8

Phenol-Chloroform
19552 - 27359
2.03 - 2.14
1.94 - 2.10
1389.2
215.4

*assessed with Agilent Femto Pulse
**Nanodrop results
The DNA fragments seem to be strongly sheared, the samples contaminated with organic compounds (especially the Magnetic bead ones) and the huge difference in concentration between Nanodrop and Qubit measurements may indicate RNA contamination.
Questions
How can DNA fragment length and purity be increased? Which are the most important measures? What life stage do you recommend using?
Ideas:

use of wide-bore pipette tips
shorten incubation time (cell lysis & proteinase K digestion)
use freshly eclosed imagines with soft exoskeleton to avoid slimy samples after cell lysis & proteinase K digestion (reason for contaminations?)
RNase A treatment

this is a fantastic question. very clear and comprehensive, informative, and using formatting effectively to drive home the point. I don't know enough to answer it though!

Welcome to Biology.SE! Strongly agree with MaximilianPress — I hope you will become a regular contributor.  ——— I don't know how easily you can obtain samples, but you may need to try all of your suggestions (e.g. do a time course for the lysis/protK step) and see what works best. In addition, my understanding is that the Nanodrop is infamously prone to inaccuracy, so I would tend to trust the other results more. You might also want to think about doing something like contour-clamped homogeneous electric field (CHEF) electrophoresis to get a idea of the size distribution of your fragments.

In absence of wide bore tips you can try cutting of some portion of the tip's front. Just ensure that the volumes you draw does not spill out. Obviously you won't be able to draw the maximum volume anymore.

Insect samples are difficult to work with primarily because they contain a bunch of polysaccharides, like chitins, that are similar enough to nucleic acids that they can co-precipitate. Standard extraction procedures (e.g. Qiagen blood/tissue kits) do not handle this well without some modifications to get rid of these contaminants. The slimy aspect of your extractions is the result of these contaminants.
Back in my day (>20 years ago) handling plant specimens, which run into similar problems with poly-phenolics, extractions were performed using modified phenol-chloroform  extractions. The specific form was a called a PCI (phenol-chloroform-isoamyl alcohol) extraction, with added B-mercaptoethanol and poly-vinylpyrrolidone in CTAB (cetyltrimethyammonium bromide) buffer.
I've had a quick search for Insect DNA extractions and found that similar protocols are in place for insects for long read-sequencing (see ref 4 in link). It looks like SDS with CTAB buffer is the way to go, followed by PC or PCI extraction.
In addition, measuring DNA and RNA concentrations by spectrometry (nano-drop etc) is unreliable - that method is great for telling you about DNA/RNA in protein solutions (the original purpose of the method), but is not really reliable enough for DNA/RNA quantitation for Gen 3 sequencing (or Gen 2 for that matter).
Instead, do as you have done, use a Qubit or other fluorescence method to quantitate and run your samples through a Bio-analyzer (Agilent) (high speed electrophoresis, with metrics on the sample added during analysis).

Thank you very much for your input!

I think @bob1's answer is good, and covers a lot of the bases.
One thing that I think is missing however is the use of a nuclear preparation as an initial step- my understanding is that this can help with a lot of the issues of phenolic/chitinous contaminants in difficult organisms like plants/fungi/insects. Here is one suggested protocol.
I would also recommend checking what's on protocols.io. Here is a protocol for insect HMW DNA prep, and here is a possibly relevant discussion.

Those might be difficult on a whole organism - generally work well from cultured cells though, and I suspect that larvae will do quite nicely, as the OP is using.


text:
What is the difference between apoptosis caused by the release of enzymes from the electron transport chain vs that from the lysosome?
I know that the mitochondria is responsible for regulating apoptosis via release of enzymes from the electron transport chain. I believe that the release of hydrolytic enzymes from a lysosome is how apoptosis occurs due to lysosomes. According to my book, the difference is that in the case of lysosomes, it leads directly to degradation. I’m just wondering how the process of apoptosis differs in each case.
Here is some info for context:
“The mitochondria are also capable of killing the cell by release of enzymes from the electron transport chain. This kickstarts a process known as apoptosis”
“Lysosomes are membrane-bound structures containing hydrolytic enzymes that are capable of breaking down many different substrates including substances ingested by endocytosis and cellular waste products... (endosomes are talked about)... The lysosomal membrane sequesters these enzymes to prevent damage to the cell. The release of these enzymes can occur in a process known as autolysis. Like mitochondria, when lysosomes released their hydrolytic enzymes, it results in apoptosis. In this case, the released enzymes directly lead to the degradation of cellular components
The reason I’m unsure if the release of hydrolytic enzymes is involved in apoptosis is because when it says “the lysosomal membrane sequesters these enzymes”, the previous text talked about how endosomes package and transport materials so I wasn’t sure if the enzymes are those materials or if it was referencing the hydrolytic enzymes.
TL;DR version: these are my two questions:

Is the release of hydrolytic enzymes from lysosomes responsible for apoptosis?

What is the difference between the processes mitochondrial and lysosomal apoptosis?


text:
Are there any eukaryotic algae that degrade EDTA?
EDTA is a frequently-used chelator in molecular biology experiments and medical applications. Buchelia-Witschel et al 2001 discuss certain bacteria have monooxygenases that are capable of degrading EDTA, so there is a basic precedent that living things can biodegrade EDTA.
The chemical properties between eukaryotes and bacteria are sometimes similar, but also frequently quite different. It isn't obvious to me that algae could not degrade EDTA, but has anyone demonstrated that there exists a species of eukaryotic algae that can break down EDTA?

From your question, I assume you're only interested in whether eukaryotic algae can degrade EDTA? "Algae" is term that describes a polyphyletic group, and cyanobacteria (blue-green algae) often get lumped in.

@acvill Thank you for the correction on the taxonomy. Yes, Eukaryotic algae only.

While I don’t have answer for this question I am curious why bacteria/algae degrading EDTA would be important.Any applications?

There may be better options for disposal, but one application might be waste disposal of a commonly-used reagent.

My own interest in this post pertains to the degradation of EDTA confounding the results of nutrient deprivation experiments.

@Galen, your comment on nutrient deprivation experiments suggests that you may be working with / considering a particular algae.  If you provide the name of your organism, you are more likely to get a targeted answer

Thank you @acvill, but this is the question I wanted to ask.

I can find no published evidence for the degradation of EDTA by eukaryotic algae. I found only one publication where algal metabolism of an aminopolycarboxylic acid was tested, specifically whether NTA could be used as a nitrogen source by different marine microalgae.1 Their results, summarized in Table VII, were that the tested organisms could not use NTA as a nitrogen source.

The possibility that NTA could serve as a nitrogen source was investigated using SS-1 medium in which NTA was substituted for the nitrogen. None of the four species of phytoplankton (Table VII) was able to use NTA as a nitrogen source. The slight stimulation shown by S. costatum at 0.073 mg/L N as NTA and I. galbana at 7.3 mg/L N as NTA does not seem to be related to NTA metabolism.

Of course, the organisms listed in the table above are just a small subset of the massive polyphyletic group that can be described as eukaryotic algae.2 Instead of directly asking whether algae degrade EDTA, we can take a naïve approach and ask do any algae encode proteins that are similar to bacterial proteins known to degrade EDTA? Section 5.2 of the review article linked by Galen 3 lists two proteobacterial strains for which the biochemical determinants of EDTA catabolism have been studied: BNC1 (Chelativorans sp. BNC1) and DSM 9103 (Chelativorans multitrophicus). For DSM 9103, a two-enzyme system consisting of a monooxygenase and an oxidoreductase was shown to be sufficient to cleave two acetyl groups from EDTA, though the monooxygenase seems to be the vital component given that the activity of the oxidoreductase could be substituted by paralogous enzyme from the NTA degradation pathway.4
Since that review, additional work has been done to characterize the catalytic mechanism of EDTA monooxygenase (EmoA) and identify structural homologs across bacteria.5 Searching the NCBI Identical Protein Group database for "EDTA monooxygenase" returns 25 hits (as of 6 July 2021) to unique proteobacterial proteins. Multiple sequence alignment of these proteins reveals a highly conserved catalytic core (red regions):

If we then find a consensus sequence from the MSA, we can use that sequence to perform a blastp search against the subset of NCBI nonredundant protein sequences corresponding to taxa annotated as eukaryotic algae by Wikipedia.

Name
Taxonomic Identifier

Mesostigmatophyceae
96475

Chlorokybophyceae
131213

Chlorophyta
3041

Rhodophyta
2763

Glaucophyta
38254

Chlorarachniophytes
29197

Euglenids
3035

Bacillariophyceae
33849

Cryptophyta
3027

Dinoflagellata
2864

Haptophyta
2830

Bolidomonas
91990

Eustigmatophyceae
5747

Phaeophyceae
2870

Chrysophyceae
2825

Raphidophyceae
38410

Synurophyceae
33859

Xanthophyceae
2833

With default blastp parameters, we get a whopping 2 hits, one to a dimethyl-sulfide monooxygenase from the endosymbiotic dinoflagellate Symbiodinium pilosum and one to a protein containing a monooxygenase-like region from Ostreococcus tauri, the smallest free-living eukaryote.6 (The same hits were obtained after blasting each bacterial protein individually to the same algal protein set.)

Description
Scientific Name
Query Cover
E value
Per. ident
Accession

dmoA
Symbiodinium pilosum
87%
1.00E-54
32.21
CAE7149325.1

luciferase-like domain-containing protein
Ostreococcus tauri
82%
1.00E-14
25.72
OUS44788.1

While both of these hits have good query coverage, the sequence identities are low, and both organisms lack (per a separate blastp) good hits to the NADH-dependent FMN reductase needed to supply the FMNH2 cofactor for the monooxygenase, though another enzyme could fill that role.
So, if algae degrade EDTA, they likely do it using enzymes that are structurally different from known EDTA-catabolizing bacterial enzymes, or by a biochemical mechanism yet to be deduced.

References

Erickson SJ, Maloney TE, Gentile JH. Effect of nitrilotriacetic acid on the growth and metabolism of estuarine phytoplankton. J Water Pollut Control Fed. 1970 Aug;42(8):Suppl:+329+.
Keeling PJ. Diversity and evolutionary history of plastids and their hosts. Am J Bot. 2004 Oct;91(10):1481-93.
Bucheli-Witschel M, Egli T. Environmental fate and microbial degradation of aminopolycarboxylic acids. FEMS Microbiol Rev. 2001 Jan;25(1):69-106.
Witschel M, Nagel S, Egli T. Identification and characterization of the two-enzyme system catalyzing oxidation of EDTA in the EDTA-degrading bacterial strain DSM 9103. J Bacteriol. 1997 Nov;179(22):6937-43.
Jun SY, Lewis KM, Youn B, Xun L, Kang C. Structural and biochemical characterization of EDTA monooxygenase and its physical interaction with a partner flavin reductase. Mol Microbiol. 2016 Jun;100(6):989-1003.
Palenik B et al. The tiny eukaryote Ostreococcus provides genomic insights into the paradox of plankton speciation. Proc Natl Acad Sci U S A. 2007 May 1;104(18):7705-10.

Fantastic work! Thank you for putting in your valuable time to search through the literature.


text:
Is centrosome one of the structures that red blood cells lose during maturation process?
Centrosomes are microtubule organising centres- their function is to facilitate cell division.
RBCs don't undergo division. They are produced via hematopoeisis in the bone marrow. It doesn't even have a nucleus, hence there is no use for centrosomes in mature RBCs and they probably don't have it.
I couldn't find any paper indicating this point. Is that true?

I was aware that mature red blood cells (RBCs) lose their nucleus, ER and Golgi apparatus. However, I wasn't sure about the centrosome! Centrosomes, being MTOCs (microtubule organising centers), have known functions beyond those in cell division. Some are cell type specific! So be wary of thinking that all cells that do not undergo mitosis lack centrosomes. I find it flawed to reason along the lines of
mature RBCs are anucleated cells =  no nucleus = no nuclear membrane = no mitosis = no mitotic spindle = no centrosomes

After a little research, I've found the following unreferenced statement in this opinion piece by Rieder, Faruki & Khodjakov from 2001 in Trends in Cell Biology:

In some cells such as megakaryocytes, myocytes and red blood cells of
higher animals, the centrosome degenerates. As a rule, however, such
cells are terminally differentiated and no longer capable of entering
the mitotic cycle.

If somebody finds a concrete example (or better yet, image!), that would be more convincing than an opinion piece.