text:
The evolution equations under different year classes of a age-structured model?
This sounds a bit complicated but I want to grab more feelings on age-structured problems. Less than 2 days to the exam so I appreciate any help.
So suppose we only get 3 year classes in a school at first. No one can repeat. If you are kicked out of school you must restart from year class $1$. No transfer-in students. Exactly $N$ freshman each year. The portion of being promoted from year class $i − 1$ to year class $i$ is given by $s_i$ (which we can again assume it's constant). Then suddenly at one year $Y$, the school changes its scheme. Only at that year the school take in $N$ year $0$ and year $1$ students respectively. Since year $Y + 1$, the school only take in $N$ year $0$ students each year.
What I want to know the most is how we should decide (1) the steady state no. of the students in year classes $1$ to $3$ before year $Y$, (2) the steady state no. of the students in year classes $0$ to $3$ since year $Y$, and (3) the school years needed to achieve the steady state since year $Y$.

I tried putting forward the Leslie matrix of the old scheme: $$\begin{bmatrix} 1 & 0 & 0 \\ s_1 & 0 & 0 \\ 0 & s_2 & 0 \end{bmatrix}$$ and found the eigenvalues would be $1$ and $0$ (with multiplicity of $2$). $\lambda = 1$ gives $$u_{1, n} + u_{2, n} + u_{3, n} = u_{1, n}(1 + s_1 + s_1 s_2).$$ Am I on the right track? Should I just plug in $u_{1, n} = N$ and then claim the steady state no. to be $N(1 + s_1 + s_1 s_2)$? Or do I need to sort of prove that?

If my thought process is correct, the Leslie matrix of the new scheme becomes $$\begin{bmatrix} 1 & 0 & 0 & 0 \\ s_0 & 0 & 0 & 0 \\ 0 & s_1 & 0 & 0 \\ 0 & 0 & s_2 & 0 \end{bmatrix}$$ Right? Then its eigenvalues are again $1$ and $0$ (multiplicity $3$), and then $\lambda = 1$ gives (let me denote the intake proportion $s_{-1} = 1$) $$u_{0, n} + u_{1, n} + u_{2, n} + u_{3, n} = u_{1, n}(1 + s_0 + s_0 s_1 + s_0 s_1 s_2) = u_{1, n}\left(\sum_{i=-1}^{2} \prod_{j = -1}^i s_j\right).$$ Did I make any mistakes so far?

I can sort of write out the year class size since year $Y$ explicitly. It should take 3 school years to achieve the steady state again right? Any less tedious and/or more persuasive ways to show the same result?

Thanks in advance.

Welcome. I have the feeling you might get a better response on crohttps://stats.stackexchange.com - we don't usually have many people answering technical stats questions like this (same goes for your other question as well).

I see. I'm actually asking if anyone can verify the general procedure I put forward instead of needing to evaluate the population with specific values.

Your results are correct if $s_i$ is the actual proportion of students which are actually promoted (and if recruitment is actually of $N(1-s_})$ so that year 1 has $N$ students including $Ns_1$ who weren't promoted). Else, if $s_i$ is a probability that applies individually, the results are different.

How is this? I added that those who are not promoted are kicked out of school. So every student is not (directly) repeated. They can of course be kicked out and come back again from year 1.

OK, hadn't seen this kicked out bit, so ignore brackets.

Still, it remains that if it is an individual probability, say rolling a dice for each of the $N$ students, you have a nonzero probability to end up e.g. with 0 students promoted (probability $(1-s_i)^N$ or all of them promoted (probability $s_i^N$).

How about if I modify $s_i$ to be the overall portion promoted from year $i-1$ to year $i$?

Yes, then your reasoning sounds fine.

Good luck with your exam!


text:
Does one wing of fly contain harmful microbes and the other contain only antibodies?
There was this study but the conclusion and it's source's relability seemed a bit off. How come a house flies one wing contain all microbe and the other don't.

In addition to @S Pr's answer, what would possibly be an evolutionary reason for flies to "care" what the microbes on their bodies do?  Seems much more likely that the presence of microbes on flies is purely accidental: they just stick to the fly's feet & mouthparts when the fly feeds.

Curiously, there is interest and research on how microbes and microbiota affect feeding, courting/sexual fitness and aggregation. Microbes also can alter social communication in Drosophila. Whether or not flies care or do anything about it is not yet abundantly clear, but there is a clear relation between microbes and Drosophila emerging in recent years. So the question is really interesting; the paper liked by OP is the exact opposite of what is interesting in the field!

I really think this question should be closed because the function of this site is not to review papers. Any question here should be about a biological problem and comprehensible on its own. The poster perpetuates the (imho) nonsense of the author by talking about fly antibodies. INSECTS DO NOT PRODUCE ANTIBODIES.

That's one of the weirdest science pubs I've ever seen, starting with reference to a 1400 year old hadith, and it's difficult to read because it's not very logical. Let's not forget that the same hadiths say that the moon and heavens followed at the same time. In reality, the moon came 9 billion years after the heavens, so a book that is wrong by 9 billion years in scientific terms shouldn't be cited as modern science facts.

Does one wing of fly contain harmful microbes and the other contain
only antibodies?

False.
A healthy microdose of skepticism is all you would ever need here. It's just a silly attempt to prove a literal interpretation of a religious text with minimal effort.
Unfortunately, the paper looks, reads, and smells like a joke. Ignoring its nonsensical title and its unintelligible sentences, it is most clearly a rather transparent attempt at indoctrination and evangelism. It is a very dirty attempt to disguise ideology as science. It does this so poorly however, that both honest laymen and honest scientists would be able to spot it within a few seconds of looking at the paper.
It is absent of properly carried out science, but it does look like a phoney mock-up of it. The "biology" contained within is completely laughable, I'm sorry to say. One example of it is the pretend experiment with an n of 1, its lack of controls, and its method section that doesn't indicate anything pertinent. Something about the use of a microscope, but an appeal to simply looking at the tubes which are photographed under some very funny backgrounds and lighting.
Personally I don't recognize the journal, it seems that it may be a fake/predatory journal, and it has been discountinued in Scopus in 2014 for whatever reason. My favorite parts? The research question:

The aim of this research was to clarify the miracle in the   hadeeth
of   the   prophet   Muhammad   (PBUH)   by studying the antimicrobial
effect of the fly wings. And to understand this aspect from the
scientific point of view.

And the grand conclusion at the end of the discussion:

Also  this  information  has  not  been  discovered  only a  few
years  ago.  this  speech  testifies  the  honesty  of  our prophet
(PBUH)band also it contain a replay to any one claiming  that  the
talks  of  prophet  are  full  of  myths moreover every word uttered
by him is right from God the almighty

I would gladly replicate this experiment for you (and the original author), it would hardly take two minutes of work (separating wings from 4 flies)... but I don't know if it would even be worth the time for any other reason than pure entertainment. This is obvious fraud and I post this answer only because the hilarity of the "paper" inspired me to. I have never stumbled onto this kind of research before, it's... perspective-building to say the least.
PS. I checked the ORCID of the author, I can find no publications that they claim to have published in journals, except for what you can find on ResearchGate. Most of the links are false links. Claims also to be a world academic champion in 2017 in microbiology (veterinary medicine), whatever that means. Claims to be a professor at Shaqra University. Can't find them on the official staff list of the university. Also, self-described:

She is the World Academic Championship 2017 in microbiology and she
was recognized as among World's 500 Most Influential Experts in
Microbiology for the Year 2017 on Earth according to International
Agency for Standards and Ratings

Hilarious!
PSS. Found a correspondence address to the author where they seems to have photoshopped some of their 'work' onto a blank journal page with formatting. I will ask the author on your behalf and let you know what they think of your question. I hope for a swift reply!

what about this paper ?  I know it sound weird for me to ask this questions but ı see a lot of papers like this. @S Pr

Nope, it's all religious zealotry. No scientific basis. I guarantee you that if i were to replicate the experiment, and falsify the results, the original authors would not accept the null hypothesis (that their religious verses are evidently falsifiable myths). That would be the honest test. Scientific papers look very different and are more rigorous. I see you are very new to scientific literature. Perhaps have a browse and see how good peer-reviewed science even looks; a simple bar chart is really not enough for any argument to hold.

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text:
T-vector creation
I'm working on TA cloning for the first time and I need to turn some existing plasmid backbones into T-vectors. I'm using a Klenow fragment (3' -> 5' exo-) to dA-tail my inserts. Is there any reason why I couldn't use this Klenow fragment to dT-tail my vectors with a similar reaction set up? Thanks in advance for any insight!

Personally I would use Taq DNA polymerase to A overhang my inserts, especially if you are PCR amplifying the insert. It leaves an A overhang on the end of all amplicons natively. All you have to do here is add some Taq at the end of the reaction and incubate for 15 min at 72 $^\circ$C, it'll work in pretty much any PCR buffer and many restriction enzyme buffers too.
Generally it is the vectors that have a T overhang, but this is a legacy of when Taq was pretty much the only polymerase around, so there is no reason you can't have an insert with a T and the vector with the A.
As a general note, often these days it is easier (and cheaper (when you factor in the cost of your time etc) + quicker) to buy a suitable TA cloning vector pre-made, they are highly reliable and should work every time, which is not necessarily something you can say about home-made ones.
Another option, if it fits your workflow, is to add restriction sites to your primers and amplify the insert by PCR. You can then digest the amplified insert and vector with the restriction enzyme of choice and get directional cloning (where your insert always goes in in the same orientation, particularly useful for protein expression). This also allows you to do things like insert short tags (e.g. FLAG, HA, V5), make frame adjustments and even modify the ends of the sequence.

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text:
Data repositories for data on the Tumor Microenvironment?
Does anyone know of any data repositories for data on the Tumor Microenvironment? I know of the The Cancer Gene Atlas and the Gene Expression Omnibus databases but I'm more curious about data on the (molecular) composition of the TME. (I'm also curious about any databases with data less specific.)

Have you seen the St Jude Children's Research Hospital Pediatric Cancer data repository PeCan?
They have a bunch of data on a wide range of different types of pediatric cancers. These are all in the form of genomes and RNA-seq. However, they also have a bunch of visualizations set up so that you can see how these profiles work from a perspective of mutations and RNA in tumours of all sorts.


text:
Are SNPs or SSR copy number variation mutations more prominent?
I'm trying to get a sense of the dominant way that mutations occur.  I have seen various numbers which seem at least at first glance to conflict, and I was curious if anyone had clarification on this.
According to Shastry, "SNP alleles in human disease and evolution" (Journal of Human Genetics 47:561–566, 2002),

In two randomly selected human genomes, 99.9% of the DNA sequence is identical. The remaining 0.1% of DNA contains sequence variations. The most common type of such variation is called a single-nucleotide polymorphism, or SNP.

However, according to Nevo, "Genetic Diversity" (Encyclopedia of Biodiversity, 2001),

Significantly, SSRs experience mutation at notably higher rates than do nonrepetitive sequences: $10^{-2}$ to $10^{-3}$ per locus, per gamete, per generation, which leads to their high polymorphism.

Additionally, according to Kashi and King, "Simple Sequence Repeats as Advantageous Mutators in Evolution", (TRENDS in Genetics 22(5):257),

Mutations that alter repeat number typically occur at rates orders of magnitude greater than single-nucleotide point mutations.

So, if variations in organisms is mostly due to SSRs, and this is mostly copy number variation, then would it be correct to say that copy number variants in SSRs is the dominant form of variation, or am I missing something?  Which one is correct, or which source is more authoritative, or is there a more up-to-date review I should be consulting?

NOTE - I added the word "mutations" to the question title to be more clear what I'm asking about.

I like @mgkrebbs answer, I think that it hits most of the high points, but I wrote a review on this subject a couple of years ago where we specifically put together estimates on the mutational load of different mutation classes (see Table 1).
Yaniv Erlich's group directly addressed the question that you are trying to answer, and they estimated that microsatellites contribute slightly more mutations per generation than substitutions:

These predictions indicate that the load of de novo STR [SSR] mutations is at least 75 mutations per generation, rivaling the load of all other known variant types.

Most estimates of diploid overall substitution mutation rates are ~50/generation, for comparison.
However, if you are interested in overall variant number you will also need to take into account an incredibly complex architecture of structural variants in basically any genome, including also plasmids, viruses, and transposons. So when you talk about the "dominant form of variation", you really do need to be a bit more specific. On a per locus or even per mutation class basis, STRs/SSRs probably are "dominant", but it's hard to argue that e.g. 1 STR mutation is more important than one transposon hop or one chromosome aneuploidy or one centromeric satellite contraction.
I'd recommend reading the papers linked for a little more context.

It's not really clear to me what you're after when you ask about how prominent or dominant these two kinds of variations are. But let's look deeper.
Guichoux et al. (1) say:

There are two main differences between SSRs and SNPs. First, SNPs are more numerous than SSRs in the genome of most species. On average, in the human genome, there is one SNP every 100–300 bp (Thorisson et al. 2005), com-pared to one SSR locus every 2–30 kb (Webster et al. 2002), depending on how SSRs are defined (Kelkar et al. 2010).This can be important for genome-wide association studies but not necessarily for other applications. Second, the mutation rate per generation differs drastically between the two marker types. SSRs have mutation rates ranging from $10^3$ to $10^4$ per locus per generation (Ellegren 2000), compared to about $10^9$ for SNPs, i.e. several orders of magnitude lower. As a consequence, SNPs are typically diallelic: In humans, < 0.1% of SNPs are triallelic (Lai 2001).In contrast, SSR loci generally have high allelic richness, often in excess of 10 alleles.

Based on these numbers, SNPs occur perhaps 80 times as often as SSRs.  Is that more prominent?
An SSR is larger than an SNP, covering perhaps one hundred to several hundred bases versus one base (by definition).  An SSR locus also occurs in multiple variants, e.g. having a copy number from a wide range, versus only two possible variants that usually occur for an SNP locus.
The mutation rates are substantially different, apparently typically 5 orders of magnitude more frequent for SSRs than for SNPs.  Note, though, that mutation rate does not directly affect the frequency of variants seen in the genomes of extant organisms: the variants present are a result of the filtering of mutations by the forces of selection.
Note also that all bases are subject to mutation, thereby producing an SNP, but SSR mutations only occur where a repetitive sequence already exists.  (Also, the likelihood of mutation becomes significant only when the existing sequence has more than a few repeats, as the mechanism of mutation depends on this.)
A main question, though, is what is the significance of the variation?  Are you interested in variation just because they mark individual genomes within a population, or are you interested in the variation of the phenotypes of individuals?  Some genetic loci are irrelevant to phenotype and some can affect the phenotype to lesser or greater degree.  SNPs can occur anywhere so can be presumed to have a "fair" chance of affecting phenotype.  On the other hand SSRs can occur where only where repetitive sequences can occur (and persist where they survive selective forces). With SSRs, varying copy number in some contexts such as protein coding sequences, may have larger phenotype effects than an SNP, and thus be subject to stronger selection forces.
My view is that both SSRs and SNPs are important and it's not terribly productive to worry about general prominence or dominance of one over the other.
Reference:
(1) Guichoux, Erwan & Lagache, Lelia & Wagner, Stefanie & Chaumeil, Philippe & Léger, Patrick & Lepais, Olivier & Lepoittevin, Camille & Malausa, Thibaut & Revardel, Emmanuelle & Salin, Franck & Petit, Rémy. (2011). Current trends in microsatellite genotyping. Molecular ecology resources. 11. 591-611. 10.1111/j.1755-0998.2011.03014.x.

The reason I'm interested is that I think that the nature and pacing of evolution is tied to the mechanism of mutation.  I think there is relevance to modeling in population genetics.  If I understand correctly, you are saying that there are far more SNPs in existence than copy-number variants of SSRs.  On a per-base-pair basis, SSRs predominate, but, since they are fewer and further between, this might not mean more actual SSR mutation instances per generation,but it sounds like it it might favor SSRs by about an order of magnitude(4 orders of magnitude less in genome vs 5 orders more likely)


text:
Nonlinear (Quadratic) selection gradients formula
Under scenarios of stabilizing or disruptive selection, we can add a quadratic component to our model of phenotype and fitness like so.

Specifically, I am not clear on where the 1/2 comes from nor do I understand the derivation of gamma, which is equal to as follows:

x is the phenotype of interest, w(x) is the fitness of said phenotypic value. uBS is the mean phenotype before selection. Any help would be greatly appreciated.

Hi, do you have a link to where those equations are from? Thanks.

Those two equations come from Lande and Arnold 1983 per the comment below this one :) Sorry!

Coefficient $1/2$ is a matter of definition/convenience. One could have written $\beta x_i + \gamma x_i^2+\bar{w}$, but then a factor of $2$ would surface in the other forumals.
The other two equations are adapted to one-dimensional case (one trait) from the Lande and Arnold's paper The measurement fo selection on correlated characters. In particular, the equation with covariance is their equation (13a), which follows from their more general equation (4) for quadratic selection. The last equation is the restatement of their (14b).
Update
Here I retake the notation of the above-mentioned article by Lande, but adopt it to the case of a single trait. Then the variance matrix $P$ is just a number (variance). Equations (14a) and (13b) define
$$\gamma=\frac{1}{P^2}C, (A)\\
C=Cov\left[w, (z-\bar{z})^2\right],
$$
which is the definition of $\gamma$ in the OP. It is essential here to use the correct expression for the covariance:
$$
Cov\left[f(z), g(z)\right] = \int \left[f(z)-\bar{f}\right]\left[g(z)-\bar{g}\right]p(z)dz=
\int f(z)g(z)p(z)dz -\bar{f}\bar{g}
$$
In our case we thus have:
$$
C = \int w(z)(z-\bar{z})^2p(z)dz - \bar{w}\overline{(z-\bar{z})^2},
$$
where $\bar{w}=1$ by definition, whereas $\overline{(z-\bar{z})^2}=P$ is the variance of the trait. We thus have
$$
C = \int w(z)(z-\bar{z})^2p(z)dz - P, (B)
$$
where
$$
p(z)=\frac{1}{\sqrt{2\pi P}}e^{-\frac{(z-\bar{z})^2}{2P}}.
$$
Let us now consider the integral
$$
\int \frac{\partial^2w(z)}{\partial z^2}p(z)dz=
\frac{\partial w(z)}{\partial z}p(z)|_{-\infty}^{+\infty} - \int \frac{\partial w(z)}{\partial z }\frac{\partial p(z)}{\partial z }dz=\\
\frac{\partial w(z)}{\partial z}p(z)|_{-\infty}^{+\infty} -
w(z)\frac{\partial p(z)}{\partial z }|_{-\infty}^{+\infty}+
\int w(z)\frac{\partial^2 p(z)}{\partial z^2 }dz
$$
The first two terms vanish, since $w(z)$ is bounded, whereas $p(z)\rightarrow 0$ as $z\rightarrow \pm \infty$.
We thus have
$$
\int \frac{\partial^2w(z)}{\partial z^2}p(z)dz=
\int w(z)\frac{\partial^2 p(z)}{\partial z^2 }dz=
\int w(z)\frac{\partial^2 }{\partial z^2 }\frac{1}{\sqrt{2\pi P}}e^{-\frac{(z-\bar{z})^2}{2P}}dz=\\
\int w(z)\left[\frac{(z-\bar{z})^2}{P^2}-\frac{1}{P}\right]\frac{1}{\sqrt{2\pi P}}e^{-\frac{(z-\bar{z})^2}{2P}}dz=
\int w(z)\left[\frac{(z-\bar{z})^2}{P^2}-\frac{1}{P}\right]p(z)dz=\\
\frac{1}{P^2}\int w(z)(z-\bar{z})^2p(z)dz - \frac{\bar{w}}{P}=\\
\frac{1}{P^2}\left[\int w(z)(z-\bar{z})^2p(z)dz - P\right]=\frac{C}{P^2}=\gamma,
$$
where we used $\bar{w}=1$, and equations (A) and (B) above.

The notation in the Lande and Arnold paper is incredibly difficult for me. If you look at 14b, could you help me understand the math that describes why the first part of 14b is equal-to the second part of 14b. I run a population genetics club and this will be incredibly helpful for us.

Essentially the same question as above in the sense that am confused about making the connection between the two equations for gamma above in this thread.

@user6817734 I have added the details of calculations. I hope it helps.

This is absolutely incredible. Thank you so very much.


text:
Using Height and Area for plotting histogram of each marker in FACS
I had a flow cytometry experiment. After normal filter with cell debris and doublet cells, I would like to gate the cells by different markers. However, I get confused to use which parameter to plot out the histogram/ scattered dot plot. More specifically, for each fluorescent channel, there are 3 options: height, width, and area. I know that width is only related to the cell size while height and area both correlate with the signal intensity. I would like to know which measurement (i.e. height or area) should I use to plot the graph?
Someone suggested that if I am analyzing different cell populations with different shapes/sizes, I should use the height. I wonder whether this is true.
Thanks!

Height is most typically used for fluorescent channels, but not necessarily for forward and side scatter. For example, you'll look at a plot and see FL2-H vs FL1-H. Experiment by substituting the different options and see how your plots change, if at all.


text:
The wood wide web strengthening/expanding the heartwood of damaged pines
My grandfather told me about a practice: girdling/ring-barking (not sure which was employed) of pine trees in order to make the wood stronger. Now, he didn't give too many details, so I had to do some research. He said the resultant, strengthened wood is called adel, adl or al, which is Norwegian.
When I searched for it, I found that these terms mean the same as heartwood, but in the context of pines. I believe that this process he explained in essence just strengthen and/or expands the heartwood of the pine. Such pine wood, that contains higher amounts of heartwood, is called ore-pine, and this Wikipedia article states that this was achieved through the removal of branches, as opposed to girdling/ring-barking, which is what my grandfather said.
Now, whether the mechanism of expansion/strengthening of the heartwood is via branching or girdling/ring-barking is one question. However, the question I am most interested in, is the mechanism of how this process happens. I know it has something to do with the increase of resinous substances in the wood, but what I'm wondering about is if this strengthening is happening via the "wood wide web". When my grandfather started talking about this, I said that it probably happened via the wood wide web. The damaged tree sends out information of its damaged state, and the other trees in its vicinity divert more resources to it, leading to the strengthening and/or expansion of its heartwood. My grandfather however, said this process did not involve the wood wide web, and that this process happened completely locally at the tree.
Which is correct (if either)?

This is a guess based on my knowledge of tree physiology. Sapwood is formed by the cambium of the tree. Perhaps when you cut off sap from the cambium by girdling, the tree cannot grow sapwood as quickly and diverts resources to the heartwood. Evidence on the role of the wood wide bed (common mycorrhizal network / CMN) in supporting stressed plants is mixed, and sometimes results in the fungi taking extra resources from the dying roots.


text:
Which fly is this?
This picture was taken in the Brazilian Amazon, upper Negro River, at daytime. The fly, about 1 cm long (maybe a little more) was flying next to us, as if to drink sweat. It finally landed on my friend's arm and I photographed it. Can anyone identify it, at least at family level?

EDIT - I've found a second pic. Left the index finger nail for size reference.

My hunch is that it's a hoverfly (family Syrphidae).

(DISCLAIMER: I'm no expert in flies and am more or less educated guessing.)

Here is an example picture of a somewhat-morphologically similar species (North American species, but whose genus is found in the neotropics):

 Toxomerus marginatus ; Credit: Ron Hemberger ; Source: Cornell University
Hoverflies are found all over the world and often demonstrate bright (often yellow) coloration. Usually, the yellow alternates with black, and there are in fact a number of bee mimics! Their sizes range quite drastically, too: 0.5-3cm [Source: Ohio State Extension].
The give-away that it's a fly (i.e., order Diptera) is the fact that it only has one pair of prominent wings. [Source: Ohio State Extension]
One of the tell-tale signs to differentiate Syrphidae from other flies is the presence of a seemingly extra vein in its wing. This is actually a false (or "spurious") vein that doesn't connect on both ends, and it parallels the 4th longitudinal vein. [Source: Britannica].

 Spurious vein (or "vena spuria"). Image source: Draw Wing

 Comstock-Needham_system. See "SV" for the spurious vein. Image Source: Wikimedia
The image in your photo does not show the wing very clearly, but I can at least convince myself something similar to the SV is somewhat visible in your specimen's left wing:

I've had many experiences of hoverflies lingering around people and following us along even as we walked through the forest for quite some distance. I think they may be attracted to bright clothing as they are very important flower pollinators.
I'm no expert in hoverflies, so my search ends here. However, if this is in fact a hoverfly, it turns out you can find online access to Thompson's (1999) "A key to the genera of the flower flies (Diptera: Syrphidae) of the Neotropical Region including descriptions of new genera and species and a glossary of taxonomic terms used" which may be of use to narrow down to a more specific taxa.
 Thompson, F.C., 1999. A key to the genera of the flower flies (Diptera: Syrphidae) of the Neotropical Region including descriptions of new genera and species and a glossary of taxonomic terms used. Contributions, Ent. International.

UPDATE:
I've actually grown fairly confident that I actually came to the correct genus in my example picture above. Toxomerus is found in North and South America, with known species in the neotropics. According to Wikipedia and inaturalist, this genus is distinguished by 1) relatively small size (6-9mm) -- check --, 2) unique abdomen markings -- check --, and 3) a posterior triangular indentation of their eye -- we can't see this in either picture.

 Eye indentation. Source: Instituto Nacional de Pesquisas da Amazônia
I'm still uncertain about species, but G.F.G. Miranda provides a list of 29 species found in the Brazilian Amazon (sourced from Macquart 1855). [Bourges & couri (2009) apparently revised these species, but I can't gain access to their publication].

 Toxomerus species found in Brazilian amazon. Source: Instituto Nacional de Pesquisas da Amazônia
Of these, non-reputable pictures of T. floralis and T. watsoni (e.g., here) seem most similar. However, due to the lack of reputability of the sources I could find (and lack of sources for most other species listed), I don't feel confident narrowing for you at this time.

 Borges, Z.M. and Couri, M.S., 2009. Revision of Toxomerus Macquart, 1855 (Diptera: Syrphidae) from Brazil with synonymic notes, identification key to the species and description of three new species. Zootaxa, 2179(1), pp.1-72.

Thank you! I've found a second pic, not sure if it helps any further.

https://www.inaturalist.org/posts/14985-identifying-toxomerus-hoverflies


text:
Which method of gene amplification for toehold switches?
My team and I are from a high school and are planning to carry out some research investigating some toehold switch riboregulators which we have designed in silico. However, we have little experience in wet lab work. We plan to order 3 genes containing an arabinose inducible promoter, a riboswitch, a leuciferase CDS and a terminator. These genes are each ~3kb in length.
We plan to order our genes from IDT, but we're not quite sure which strategy to deploy to amplify these genes before adding them to a cell free extract for characterisation.
The four methods we have narrowed it down to are as follows:

Order the genes in plasmids, linearise, PCR,  ligate, add to cell
free extract for characterisation.
Order genes in plasmids, transform straight into E. coli, culture, mini prep, add to cell free extract for characterisation.
Order genes in plasmids, linearise, PCR, ligate, transform into E. coli, culture, mini prep, add to cell free extract for characterisation.
Order genes as gBlocks (not in plasmids), amplify with PCR, ligate into plasmid backbone, transform, culture, mini prep, add to cell free extract for characterisation.

We'd ideally like to use the method which is most simple as we will not have a lot of time in the lab, and we don't have much experience.
Any advice on choosing between these strategies would be much appreciated!

The strategies you're thinking about seem fairly general, rather than specific to toehold switches. Can you clarify if there is something in particular that you're concerned about regarding toehold switches, or is this a more general question about assembly strategies?

@jakebeal There isn't anything really which toehold switches will affect. So this is more of a general question about assembly strategies, yes.

Welcome to the Biology Stackexchange! Are all three plasmids going to have the same Promoter, CDS and terminator, but a different riboswitch?

I basically agree with Bob, but wanted to add a few more details.
Out of the options that you have available, option 4 is the most common way to do it. I wouldn't consider options 1 and 3. I don't think option 1 will really work to produce the amount of plasmid that you need, option 2 is a much easier way of amplifying it. As for option 3, you shouldn't need to amplify the plasmid before transforming it, on the IDT website they say:

All of our synthetic gene products are provided in an optimized
cloning vector that is ready to be transformed into E. coli.

Between options 2 and 4, it comes down to time and cost. Unless you have a deal for free synthesis from IDT, the cost of option 2 is going to be around £1100 vs £330 for the gBLOCK (although another consideration here is that the maximum gBlock length listed on the website is 3kb). In terms of timing, the website says it will take 5-8 business days before shipping a gBlock vs 25 business days for the plasmid. Someone who is an experienced cloner would definitely be able to clone the gBlock into a vector faster than that (might take as little as 1-2 days), but if you're first-timers you might find that it takes you months to get it to work (the first time I cloned it took me a long time).
Another factor for both options 2 and 4, depending on how you do it, is that if you're using different toehold switches then you'll either have to order 3 plasmids/gBlocks with essentially all the same stuff apart from the toehold switch, or replace the toehold switch using something like an inverse PCR.
Assuming you are an iGEM team, another option, although maybe not the best if you have free DNA synthesis from IDT, is that the iGEM distribution kit  will be distributed around the end of June. This kit should contain samples of the luciferase CDS which you can amplify using PCR with primers to assemble into a vector of your choice. I would only recommend this method if you're limited on synthesis capability/are having issues with amplifying your gBlocks. You also wouldn't have the promoter/terminator/toehold switch, so you'd need to include them.
To summarise:

The best way (if you have the funds) is option 2
A slightly cheaper way is to order the full plasmid for one toehold switch and then use PCR to replace the toehold switch for the two others
The 'normal' way that I in my experience is the way most people do it is option 4, this is also the quickest if you know what you're doing
The cheapest way to do it is using the iGEM distribution kit

In my experience none of the options are the most common, though 2 would be the closest. Generally most people I know would do just what I describe in my second paragraph - it's quick, cheap and easy. Should be able to get the primers for this for <$100.

That's interesting. Although everyone always talks about "getting rid of cloning", I haven't encountered a situation yet where an academic lab has used option 2, for the reasons that I outlined above. Any time I have done cloning it has been by amplifying a gBlock and either using Gibson assembly or restriction digestion to insert it into a vector of choice (option 4). Perhaps it varies from field to field.
I agree that already having the gene in a plasmid and amplifying it out is the easiest way, but as OP mentions they're at a high school they're unlikely to have it lying around.

My whole academic career (~20 y) I've been amplifying sequences from various genes and then inserting them into plasmids for gene/protein expression. Usually by including restriction sites on the primers. gBlocks etc are relatively recent, and still much much more expensive than a pair of primers, a couple of common REs and a PCR. Synthetic DNA of lengths over about 200 bp have only really become cheap enough to be feasible in the past 7-10 years. Smaller genes are cheaper in plasmids - I can easily get $0.19 per bp gene in plasmid for genes <8 kb = ~$600 (425 GBP) for 3 kb.

Option 2 is your simplest and easiest. It should give you unlimited plasmid to work from. Note that ordering pre-made sequences of this size isn't particularly cheap and might take a few weeks to get to your destination.
It may be easier (quicker + cheaper) to find a plasmid with the luciferase CDS and terminator already built in. You could then amplify this with primers incorporating your promoter and riboswitch sequences and some restriction enzyme sites to generate a product that looks like this:
RE1-ara-riboswitch-luciferase-RE2
You can then clone this into a(nother) plasmid cut with RE1 and RE2.
Another option is that if you have a plasmid with a luciferase gene in it already and a multiple cloning site (MCS) you can order complementary primers (note single-stranded DNA, anything less than about 150 bases is fine) containing the araC and riboswitch sequences flanked by two restriction sites, you then anneal the two primers, digest off the ends and ligate. You just have to be careful with making sure that everything is in-frame. I don't know anything about riboswitches though so frame may not matter so much.
As you have limited time I suggest buying chemically competent cells for heat-shock transformation (if you don't have some already); make sure you don't get electro-competent ones, as these will only work with an electroporator.  Also get a commercial mini-prep kit (e.g. Qiagen plasmid mini-prep). These are easy to use and reliable.
I think in option 1 you have something a bit odd going on - why would you digest your plasmid then re-ligate it, when it is already in the form that you are going to use?
I am not quite sure why in options 3 and 4 that you think that these are any different in terms of the end result to option 2.


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Why do you need to calculate the lane normalisation factor when doing western blot data analysis?
I am learning about western blot data quantification from some online resources. I have read about methods to normalise the data.
I have seen in a number of resources, such as this handbook and this video when normalising western blot data, they use a lane normalisation factor. For example, if using a total protein stain to normalise the integrated density of your target protein bands, they first obtain the integrated density of all the lanes stained with the total protein stain. Then they find the lane with the highest integrated density value. For each lane, its integrated density value is divided by the highest integrated density value - the resulting value being the normalisation factor of the lane.
Then when doing the normalisation, for each lane, the integrated density of the protein band of interest is divided by the lane normalisation factor.
However, I am not sure what is the reasoning behind calculating the lane normalisation factor. Why can you not just divide the integrated density of your protein band of interest by the integrated density of the total protein stain region for each lane?
Any insights are appreciated.

Read point 2 of your handbook: II.TheIdealWorldvsTheRealWorld.

Thank you, I think I understand now - it's to correct for variations in total protein signal between lanes.


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Can you use images taken at different exposure times in western blot image analysis?
I am doing Western blot data analysis where I have images from a number of experiments (where the samples in the experiments are biological/technical replicates). For each experiment, I labelled the membrane with antibodies against the same proteins of interest.
I want to combine the data from these experiments so that they are all displayed on the same graph.
I took my images at a variety of exposure times (e.g. 5 sec, 30 sec, 1 min, etc). For one protein X, I have found that the images taken at 30 seconds exposure are the best for analysis, as the bands are saturated at higher exposure times.
However, for one experiment, for the protein X labelling, I cannot use the 30 second exposure image for my analysis as there are ghost bands. I have an image taken at 5 seconds exposure which I can use instead.
I was wondering when combining data from western blot images from different experiments, for a specific protein of interest (e.g. protein X), do all the images have to be taken at the same exposure time? All the protein X images I will be using were taken at 30 seconds exposure except for one which is taken at 5 seconds exposure.
Any insights are appreciated.

Generally no, you can't directly compare them and you can't easily compare across gels. I have done many hundreds of western blots, a large proportion of which were for quantitation, and in general quantitation on western blots is difficult (because of inconsistency run to run) and not highly quantitative - a 2-fold change is detectable and probably real, but a 10% change - unlikely to be real.
If you want a real example of how to compare and observe this (works with the Bio-Rad system or others where the transfer is in the same space) - run exactly the same amount of protein on two identical gels, place them both into the same transfer tank. Compare protein stain on the membranes after transfer. The one closest to the black terminal will have stronger staining = better transfer.
However, what you can do is convert the numbers you get into a relative expression on each of the blots and then use those numbers for comparison.
For example you compare expression of protein X to a normalizer (often B-Actin) and get fold-change data out of that, or you can compare Sample X untreated to treatments A, B, C and get data out that way.

Thank you for your reply. Just to clarify, I can compare the images at different exposures, but only after normalizing the target protein signal to an internal loading control?

Correct. In fact you need to do this for every blot that you run. You cannot pool results from different membranes otherwise, even if all the conditions under which they were run and probed are the same.


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What are evolutionary reasons for humans' limited regeneration abilities?
As far as I understand (I am not a biologist), a ability of a species arises during the evolution if:

It increases reproduction chances
It is not too energy expensive
It is physically possible

At the same time, if a human loses his/her hand/eye/tooth/almost anything, it won't regrow. However, if such regeneration was possible

It would definitely increase reproduction chances, because it would make it much easier to survive (or maybe something even more direct, like reproduction of sexual organs)
Ability to regenerate isn't supposed to require any energy until the moment such regeneration is needed (and, therefore, energy spending is justified)
It seem to be indeed physically possible, at least for some organs (Wikipedia provides examples of species who can regenerate their teeth, body parts (tails) or even lungs)

This seem to be strong reasons why this ability should have been gained during the evolution, but this didn't happen. Why?

I think the claim #2 (energy) requires more detailed examination of what regeneration involves. Also, the ability to regenerate is rather an exception than a rule among animals, so singling out humans is rather anthropocentric.

In order to understand limits of evolution of regenerative abilities you must understand the molecular biology of embriogensis including the main genetic players.
https://journals.biologists.com/dev/article/147/14/dev190629/224370/Mechanisms-of-human-embryo-development-from-cell
You should appreciate the complexity of embriogensis! Obviously, this puts enormous constraints on evolution.


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Pathological mechanisms in asthma
What is known about the precise molecular mechanisms involved in asthma?  For instance, is it known why the bronchi tissue alteration cannot be reversed?  Were any new mechanisms discovered in the last decade or so?

I don't know the answer myself off-hand, but this seems like a rather basic question, so you should probably look it up somewhere else (such as Wikipedia, or a medical textbook) rather than asking people to explain it here.  On this site you're only supposed to ask for information that you couldn't easily get ready-made.  If you have in fact looked it up already without success, you'll get a better reception if you say so and tell them something about where you've already looked.

Yes I understand, but my hope was that someone who knows well that topic could give me the great lines, so that I could find by myself the details more easily.


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Exclusion and inclusion criteria for healthy controls in colorectal cancer study
I'm working on colorectal cancer, and I'm facing a problem in the sampling step, I can't figure out how to choose healthy subjects. In the literature, I've found many criteria that are different and sometimes not clear or the author don't mention how he chooses his control samples.
PS: I'll collect stool samples, not tissues.

Welcome to the world of poorly designed studies. Not saying yours is or will be, just  saying the confusion you feel isn't unwarranted. I wish I could offer you a solution.

https://medicalsciences.stackexchange.com

Providing more details (e.g. what you are going to study, how you are planning to do it, how others have done it) would invite good answers to this question. Also I agree with @anongoodnurse: selecting controls for a case-control study can be challenging. (There is a nice section on this in a textbook called  'Gordis Epidemiology', that might help. https://books.google.co.in/books?id=Xwl0DwAAQBAJ)


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Identify orange-brown, teardrop-shaped insect in central Europe that was found indoors
TL;DR Can someone identify the species of the insect in below picture? Editing of the wording of this question (if possible) to make it easier to find by using search engines is welcome.
I caught this insect and was wondering what species it was:

Here a milimeter-scale and an USB-plug next to it to allow to estimate the size (sorry that they are so unsharp but I hope it is sufficient to get the size):

I was not able to find it by reverse image search or by searching for some keywords using google. I also checked some questions here but did not find any that seems to be about such an insect.
Following description/keywords are added to help someone searching for such an insect to find this using word-search. If someone can formulate this more search-engine-friendly to make this more likely to be found, please edit or add a comment so I can edit it:

Teardrop-shaped
six legs / 6 legs
two antennas / 2 antennas
orange-brown / brown-orange
size about 3mm / 3 millimeters / 0.11811 in / 0.11811 inches (I hope my calculation from mm to in is correct)
if touched by glas or paper, it remained motionless for about 1 to 2 minutes
no wings / not airbourne / walking
able to walk up walls
it was found indoors
the region is central Europe

Many Thanks in Advance.

For everyone who may be waiting for an answer to this or who at some point in the future find this through google: I have found the species fo the respective insect, it is Gibbium psylloides (https://en.wikipedia.org/wiki/Gibbium_psylloides).

Thank you for posting an answer to your own question. Please note that you are much more likely to get a positive response to your answer if you explain why your identification is correct. Specifically, please [edit] your answer to add discussion of key features that led you to this conclusion and supporting references or at least validated images. Without this your answer is indistinguishable from opinion. ——— Please consult the [help] pages for additional advice on [Answer] effectively on this site.

I think it may have been a tick of some kind, one that had already fed and was round (hungry ticks are sort of flat).  I couldn't say exactly what species.
I'm not sure, but that's the only thing I can think of that matches this photo.  I'm no professional, so another person here may be able to do better.
Ticks have eight legs (they're vaguely related to mites, and, even more distantly, to spiders), so I was puzzled at first by you saying it had six legs, but possibly what you thought were its antennae were its other pair of legs?  (Ticks have no antennae).
https://en.wikipedia.org/wiki/Tick

Thanks for the answer. I see the similarity with tigs but I doubt this is a tick. For onc, I think it really had antennae because it moved them while walking instead of using them as legs. In addition, ticks would likely not remain motionless if touched because they have to grab prey walking past them when sitting in the gras. Also, ticks have their legs somehow streched out to the side, this one has them kind of V-shaped. I agree that from the general description this could be a tick so this answer may be good for someone else finding this through google. Please note that I did NOT downvote.

Fair enough!  Thanks for the comments, that makes sense.


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I just do not understand how myelination speeds up action potentials
first off, I've watched like 7 youtube videos, read a bunch of articles, and the explanations for why myelination actually increases action potential propagation differ each time are very vague. I've read and heard that because action potentials only occur on the nodes of Ranvier, that it simply takes less time for them to travel down an axon. I get that, but I was under the impression that increased +ve charge = further generation of action potentials. if that is the case, wouldn't that mean that myelination should slow it down as there is less +ve charge influx? I understand that it takes time for action potentials to be generated, but is the variable affecting the rate at which they occur not the influx of +ve charge, and wouldn't this positive charge be less if there are fewer action potentials generated? it's not like myelination decreases the time taken for particular na+ channels to open, right? ftr, I'm not doubting that myelination speeds up action potential propagation, I just don't get it and the explanations are contradictory (nothing that I've mentioned here is contradictory to my knowledge FYI, I just mean that I have heard other contradictory explanations). I understand that it insulates the neuron, thus preventing k+ leakage. Does the insulation possibly increase the rate of propagation by decreasing the ability of extracellular ions from interacting with intracellular neurons? I would understand that I suppose since then there wouldn't be positively charged particles on the exterior exerting electric force upon the intracellular +vely charged particles, thus making it easier for +ve charge to spread throughout the negatively charged axon. even if there is any truth in that, could that possibly compensate for the decrease in intracellular sodium that the decreased amount of na+ influx channels would imply?
thanks to anyone who responds! a long one for sure, lol.

Hi @William. Good question. Would you care to edit your post for legibility? It may help with getting more people to help you out with an answer! At the moment it's a verbose and hefty chonk of text, I'm sure you'll agree!

I think this is a duplicate; if you're still not following then maybe you can distill it down to some more specific uncertainty, but I'd recommend dropping the things you think you know and trying to relearn how it works from that linked Q&A.


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By what mechanism do cephalopods regrow limbs?
As far as I can tell, there are two types of limb regeneration: epimorphosis and morphallaxis. Epimorphosis is used by animals such as amphibians and various phyla of worms, while morphallaxis is used by hydras.
I have been trying to find out which of these mechanisms is used by cephalopods, and no reliable source paints a clear picture one way or the other. Can anyone here help?

well gastropod mollusks use Epimorphosis, so it is likely cephalopods use the same mechanisms. Basically all triploblastic organisms use this mechanism so phylogenic bracketing points towards it as well.
morphallaxis is generally restricted to very simple organisms.
Regeneration of the visual system in gastropods (Mollusca)
Oksana Tuchina  Victor Benno Meyer‐Rochow
Neural regeneration in gastropod molluscs
Author links open overlay panelStacia B.Moffett
Regeneration of the visual system in gastropods (Mollusca)
Oksana Tuchina  Victor Benno Meyer‐Rochow (alternative link)


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Does transcription terminate at the end of antibiotic resistance genes on the Duet expression plasmids?
I am working with the pETDuet and pRSFDuet plasmids to express RNAs under control of the T7 promoter. On these plasmids the antibiotic resistance genes (Amp for pET, and Kan for pRSF) are downstream of the T7 promoter site where I cloned in my genes. The Amp and Kan genes are transcribed in the opposite orientation of the T7 promoter and since these genes are downstream from where I inserted my genes, readthrough transcription from the resistance genes would result in a sequence complementary to the RNA sequence I wish to produce. My question is: do the antibiotic resistance genes in these plasmids have a built in transcription termination mechanism?
I have used the ARNold tool: http://rssf.i2bc.paris-saclay.fr/toolbox/arnold/index.php to look for Rho-independent terminators at the 3' end of the antibiotic resistance genes and in the region downstream. There was one hit for the pETDuet plasmid but nothing for the pRSFDuet plasmid. There were no hits for the  pCDFDuet or pCOLADuet plasmids either. My sense is hairpin terminators were not specifically incorporated to stop transcription downstream of the antibiotic resistance genes. The other possibility is the antibiotic resistance genes terminate in a Rho-dependent manner, is this known to occur for the Amp and Kan genes?

Generally they do, however this is often doesn't completely stop transcription run-through.
Note that in a T-7 reaction there shouldn't be any transcription of the Amp or Kan genes as these are run off promoters that  the T7 phage RNA polymerase won't be able to transcribe off as it is extremely specific for the T7 promoter.
The solution if you are really worried is to linearize your plasmid immediately downstream of your terminator but before the AMPR gene using a restriction enzyme with a single cut. This will also give you a specific size sequence being transcribed, which is very useful if you are determining copy number in say a qPCR.


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When analysing phosphoproteins via Western blot, why is total protein level of the target protein recommended as an internal loading control?
I am analysing the expression of a protein kinase via Western blots, and it is a phosphoprotein. I have labelled my membrane with antibodies against the phosphorylated form of the protein (using phospho-specific antibodies) and also for total protein (using pan-specific antibodies).
I have read that when studying phosphoproteins, you should normalise the phosphorylated protein signal to the signal for total protein. In this journal paper, it states that the Journal of Biological Chemistry recommends that “signals obtained using antibodies specific for phosphorylated epitopes should be normalized to the total protein level of the target protein”.
I have seen in other journal papers analysing the expression of a phosphoprotein via Western blots that they also normalise the phosphoprotein signal to total protein signal (of the target protein).
However, I am not sure why the total protein level of the target protein can be used as an internal loading control?
I have read in this handbook that internal loading controls are endogenous reference protein(s) that are present in all samples at a stable level and unaffected by the experimental conditions.
I want to compare the expression of my phosphoprotein between samples of different experimental conditions. I know that normally total protein stains such as Ponceau S or housekeeping proteins are used as an internal loading control and for normalisation.
But why is it recommended that you should normalise to total target protein (when looking at phosphoproteins), because one does not know whether the expression of total target protein will be stable between different experimental conditions? Any insights are appreciated.

It is because you don't know how your treatments affect the expression of the total specific protein; perhaps it down- or up-regulates the amount of protein being made, but the relative proportion of phosphorylation doesn't change. For example:
Imagine you have 4 samples from different treatments of which you have loaded an equal amount of total protein (lets say 50 ug total, note that this is too much to load on a WB for accurate quantitation; usually you want less than 10 ug). You run a western blot and find the following numbers (completely made up BTW) for the specific protein. In this scenario, 1 is untreated control.

10 ug
20 ug
10 ug
20 ug

You then try the phospho-specific antibody and get the following

1 ug
2 ug
3 ug
1 ug

What does that tell us about the 4 treatments?
Well, it tells us that 2 is up-regulating the total protein, but the relative proportion of phosphorylation is the same. In 3 phosphorylated protein is up-regulated (3x) but not total protein and in 4 total protein is up-regulated but phosphorylated is down-regulated (0.5x).


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In phospho/pan analysis in Western blots, what is best way to normalise to an internal loading control?
I am analysing the expression of a protein kinase X that is a phosphoprotein through Western blots. I have labelled the membrane for both the phosphorylated form of the protein and also for the total target protein. I am comparing the expression of my phosphorylated protein between different experimental conditions.
For each sample, I have calculated the proportion of protein that is phosphorylated by dividing the phosphorylated protein signal by the total target protein signal. I have also done this to ensure that the differences in phosphorylated protein between samples is not due to variations in total protein levels under different experimental conditions.
However, I want to normalise this ratio to an internal loading control, since I think that it would be incorrect to assume that the expression of the total target protein is uniform between the different experimental conditions. I have read when comparing the expression of a protein amongst different experimental conditions, you need to use an internal loading control.
I have used a total protein stain (Ponceau S) instead of a housekeeping protein as my internal loading control, since studies have shown that expression of housekeeping proteins can vary between different experimental conditions and that the level of total protein between different experimental conditions is more consistent.
I am currently debating between the following two methods of normalisation:
(1) (phosphorylated protein X/Ponceau)/(total protein X/Ponceau)
(2) ((phosphorylated protein X)/(total protein X))/Ponceau
I think that method (1) is the more correct way to normalise the phospho/total protein X ratio, since you would want to obtain the relative abundance (to Ponceau) of both phosphorylated protein X and total protein X before calculating the phospho/total protein X ratio.
However, I would was wondering if anyone knows whether this reasoning is correct? Any insights are appreciated.


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Motion of Fallopian tube fimbriae
The fimbriae at the ovary end of Fallopian tubes are described as "sweeping" eggs from ovaries into the tubes:

When ovulation is about to occur, the sex hormones activate the fimbriae, causing them to swell with blood and hit the ovary in a gentle, sweeping motion. An oocyte is released from the ovary into the peritoneal cavity and the cilia of the fimbriae sweep the ovum into the Fallopian tube.

Is there more detail known about the "gentle, sweeping motion" of the fimbriae?  For example, is it a single swelling "sweep" per cycle?  Is the motion of the fimbriae against the ovary isolated from or accentuated by other movement of the body?


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Does SARS-COV-2 produce toxins?
I wonder whether toxins are produced only by bacteria and fungi and not by viruses. Looking at the paper below, I read that several toxic proteins are produced by SARS-COV-2. Can these proteins be considered toxins?
Reference:
Lee, JG., Huang, W., Lee, H. et al. Characterization of SARS-CoV-2 proteins reveals Orf6 pathogenicity, subcellular localization, host interactions and attenuation by Selinexor. Cell Biosci 11, 58 (2021). https://doi.org/10.1186/s13578-021-00568-7

Please define what you understand by the word “toxin”. (And if you do that you will presumably be able to answer the question yourself.) As far as I am aware, the words has no unique scientific meaning, outside, perhaps, the specialism of bacteriology. It is enough that viruses infect and often destroy cells without getting into semantic arguments about toxicity.

It seems that the key here is the definition of "toxin" in a biology context. The best definition I have found is this one from Merriam-Webster:

Definition of toxin
: a poisonous substance that is a specific product of the metabolic
activities of a living organism and is usually very unstable, notably
toxic when introduced into the tissues, and typically capable of
inducing antibody formation.

Key aspects of this definition include that it is "a specific product of the metabolic activities of a living organism " (so e.g. cyanide or even HCl would not be considered toxins) and that it is "notably toxic". It's the latter characteristic that the SARS-CoV-2 proteins fail, as your article link shows at most roughly 50% cell mortality when the various ORFs are overexpressed in target cells.
Basically, those proteins are nowhere near toxic enough to be considered "toxins".
A WHO publication on biosafety (Lab biosecurity guidelines, Sept, 2006 WHO/CDS/EPR/2006.6) includes the description:

Toxins  are  poisonous  substances  produced  by  living cells or
organisms.

And you go to an English dictionary to find the meaning of a supposedly scientific word?

As I said, it's the best definition I could find. I think the "specific product of a living organism" and "notably toxic" aspects are key. I did not check dictionaries in other languages, but as you imply, those may provide additional useful information :)

I am not interested in dictionaries in any language unless I wish to find the first recorded usage of a term. For scientific meaning I would either consult a document on standards or nomenclature in the area or research usage in scientific publications.

Thanks. I was going on my own usage as a practitioner and finding a pre-existing definition that matched that.


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Which trophic level has the highest population in a food chain?
The basic trophic levels in any food chain are the producers, the consumers and the decomposers. Most of the websites I've searched say they are the decomposers. But shouldn't it be the producers? In any given food chain, the number of individuals progressively decreases towards the higher trophic levels.

The concept of a trophic "level" really only applies within the category of consumers (and even there it's only a rough approximation).
This is because some consumers eat producers (e.g., herbivores eating plants), while others eat other consumers (e.g., carnivores).  So the notion of a "level" is basically counting how far something is from producers, which in turn creates the pyramid structure of bigger and badder predators eating smaller predators, and so on. This is really very specific to predatory animals because it is only in the context of predation that size goes up and numbers go down as photosynthesis becomes more remote.
With other types of interactions like parasitic worms infesting your gut or mosquitos sucking your blood, you could categorize the organism as a consumer and count it as another level removed from producers, but since it's only tapping a small amount of resources, rather than the more total interaction of predation, the ecological consequences are very different, and you can have a lot more mosquitos than people in a way that you can't have more wolves than deer.
So it is with decomposers as well, for yet another different reason. Decomposers don't need to be big (though they can be), since they aren't trying to hunt and kill something, they're extracting value from something that is already either dead or cast off. Thus, once again the concept of "level" doesn't really apply, and a vast number of flies can be born from the corpse of a single dead lion.


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Fluorescent purple flowers?
Do some flowers fluoresce? Some deep purple Lobelias and larkspurs seem to glow and confuse my eyes in sunlight. It reminds me of those blue LED Christmas lights (they come in strings with other colors) - the blue ones only seem to be showing two colors at once.

Many flowers fluoresce under the influence of UV and white light. The compounds that fluoresce are called Betalains, and are either red/purple (betacyanins) or yellow (betaxanthins).
It is thought that this is a method of making the flowers more obvious to insect pollinators. However, there is some debate about whether the fluorescence contributes to this much compared to just the bright colours.

Fluorescence is a way of making brighter colors, though.  Ever looked at day-glow colors under UV light?  It's also common to fluorescent chemicals ("optical brighteners") in laundry detergents, in order to make colors (especially whites) appear brighter: https://en.wikipedia.org/wiki/Optical_brightener

@jamesqf - that's exactly the point of the papers linked in my second paragraph. It isn't known how much the fluorescence contributes to attractant for insects as the colours are already bright in the ranges that insects use. Fluorescence always puts out less energy than is put in because of the energy cost of the electron excitation step, so while those colours may be bright under UV alone, with the additional very bright white light spectrum, the fluorescence is somewhat lost.

We may be thinking about different things.  I'm not thinking of UV colors that are visible to some insects & birds, but of fluorescence absorbing the incident UV light and re-emitting some of the energy in the human-visible range.  Just as for instance day-glo orange is orange under inside lights (no UV), emits nearly the same shade of orange when you illuminate it with a UV light in the dark, and is really bright orange in sunlight because both oranges combine.

@jamesqf, no, that's what I am talking about too. The evidence for significantly brighter signals for plants emitting in the visible spectrum isn't there. It looks like <10% increase in brightness.


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Have orcas ever been observed to be eaten by another species?
Orcas are well known to prey on just about everything that lives in the ocean, even including great white sharks. Yet in a lot of cases I know that the food chain is not unidirectional, there are many cases where even so-called apex predators will get attacked and eaten if they are weak and let their guard down (the eternal rivalry between lions and spotted hyenas comes to mind).
My question is this, are there any known cases of orcas being killed and eaten by another non-orca marine mammal (i.e., not humans)? Such as a lone adult without a pod being attacked, or a calf being taken when it's mother wasn't looking?

Since orcas are difficult to observe, such an event has likely not been observed even if it happens. For example, members of even relatively well-observed pods like those in the Salish Sea area (Pacific Northwest) are almost never observed to die. Rather, they are deemed dead after not being observed for perhaps a year or two.

This Orca Network births and deaths page may be useful: https://www.orcanetwork.org/Main/index.php?categories_file=Births%20and%20Deaths

No, there are no known cases (unless anecdotal evidence from lifelong marine mammal observers exists, which is possible).
The best candidate for such a phenomenon would be the sperm whale due to its large size and the presence of teeth. Baleen whales could potentially injure an orca, but would not be able to eat it. However, I think even in the case of sperm whales, killing and eating an orca would be very improbable for a couple of reasons.
1 teeth size and function sperm whales have external teeth only in the lower jaw, and generally swallow prey whole 1. They cannot effectively grasp or tear prey, which would make it very difficult for them to eat an orca even if they wanted to.
2 previous accounts of interactions Accounts of interactions between sperm whales and orcas suggest that orcas are not scared of sperm whales, and that sperm whales avoid orcas when possible and engage only in defensive behaviors during interactions. This article in pictures documents an interaction between orcas and multiple male sperm whales in which the sperm whales are successful at avoiding the orcas, but do not attack them. Playback experiments such as this one have noted that playing orca calls cause sperm whales to stop foraging efforts and spend more time at the surface, again suggesting that sperm whales are more likely to try and get away from orcas than they are to ever willingly attack them.


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Do mutations occur while growing virus for preparing inactivated viral vaccine?
The development of mutations in virus is reported to happen during replication, especially for an mRNA type virus like SARS-COV-2

Viruses that encode their genome in RNA, such as SARS-CoV-2, HIV and
influenza, tend to pick up mutations quickly as they are copied inside
their hosts, because enzymes that copy RNA are prone to making errors

My question is do mutations occur while mass manufacturing inactivated viral vaccines?
Simply because the process of manufacturing inactivated vaccines replicate the virus in large quantities, there might be large number of mutations occurring. Will this make each batch different from one another? Or do they control the replication process to limit the mutations?
Veritaisum did a recent video on the study on mutations in long term evolution experiment of E.Coli. It is clear that mutations do occur even in standard controlled environment.

I already asked in medical sciences stackexchage https://medicalsciences.stackexchange.com/q/27469/22302. Reposting here for visibility

"inactivated viral vaccines" - note that the major Covid vaccines are not of this type. Moderna and Pfizer are mRNA vaccines. Astra-Zeneca and Janssen uses harmless non-Corona viruses.

Please don't repost questions. For this one, this site is more appropriate than Medical Sciences.

@MSalters there is a significant portion of covid vaccines manufactured using inactivated viruses. And mRNA vaccine is not a viable in many parts of the world. Hence asked this question

@MattDMo I was tempted to repost the question here as I did not get answers in the other portal. No harm intended; just opening up to wider audience and seeing if I can find answers. I could not get a solid result while googling too.

@karthikeyan I'm sure there was no harm intended, but it's against the Stack Exchange rules. If you didn't get a result elsewhere after a few days, just delete the question and then repost it here. We just don't want multiple parallel discussions going on at different sites.

Very simply, mutations do occur, as they do for any cultured organism. This is a well recognized problem in many fields of biology where organisms are cultured and remains in particular a problem for cultured mammalian cell research.
As far as I know there is no method for slowing or altering the rate of mutation as this is an inherent part of the RNA-dependent RNA polymerase. This happens irrespective of the environment the virus is grown in and is one of the ways that viruses of this sort combat the immune system and ecosystem changes - they produce so many mutations over the generations of viral replication that some of the progeny will be more fit in the environment in which they are replicating.
In fact, they produce so many mutations at each replication cycle that these viruses can not be called a single "species" they are actually what is referred to as a quasispecies.
It is part of the difficulty of producing vaccines against things like influenza and related viruses as they can change from the phenotype seen in the wild. However, we have methods of checking that the viruses are the same or similar enough antigenically that it makes no difference for the immune response to the vaccine.

IIRC, you can change the rate of mutation by changing codons. There are 64 codons for 20 amino acids, so not all mutations will change the resulting virus. For example, "CUA"  and "UUA"  both encode for Leukin, but "CUA" is more stable. That's because an A->U mutation would produce "CUU" and "UUU", but only CUU encodes for Leukin.

@MSalters - yes, this potentially outputs stable functional viruses, however in a normal infection (in the lab) defective particles outweigh the functional by some orders of magnitude. The error rate of the RdRP is constant for the virus, so even if you have the CUA codon, it doesn't mean it will change the rate at which it is mutated, just that functional virions will more likely have this codon.

@bob1 I haven't looked at this in years, but I believe the codon can actually affect the mutation rate and certain codons are more prone to slippage and are more likely to be read by the wrong tRNA than others. Take this with a huge grain of salt, I am vaguely remembering conversations with colleagues from several years ago, but I do think there is something along those lines.

@terdon - in what systems? We are talking about an RdRP in viruses here.

@bob1 huh, good point. If (big if) my memory serves me and there are indeed different rates of error for different codons, I would expect that to also be the case for viral RdRP. But I'm really just making wild guesses and assumptions based on vague recollections so I certainly won't press the point.


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Can insects be killed by intense visible light?
Some mosquitoes and flies were sitting on the white ceiling. I took a very powerful flashlight to try and daze and then vacuum them.
When I put the flashlight's head against the ceiling (with a mosquito between the light and the ceiling), I heard buzzing for two seconds... then it stopped. I removed the light, and, to my utter amazement, the mosquito was dead! Tried it several times on other small flying insects (including medium-sized flies) and not a single one of them could resist my light saber! :)
Could they really be killed by intense visible light (concentrated and reflected by the ceiling)?.. Note that the flashlight is not getting hot so it wasn't about heat (I can put it against my arm and it feels just barely warm).
EDIT
I would have to disagree with the comments.

First, why the downvotes?.. :)
NO, the insects are not being crushed by the torch. There's a lot of space between the torch's crown and the ceiling. As a proof — small mosquitoes and flies are quickly dead but a big fat disgusting fly was just buzzing around and then flew away!
NO, it's not the intense heat that kills them... at least not in contact form. Like I said, I can easily put the torch's head to my palm and it feels barely lukewarm.
Sentences like "skin can't be broken by light from a torch" or "insects are flying around in full sun and don't drop dead" are not substantiated, they're just some random opinion from Internet. For the contrast, here's a scientific article: https://doi.org/10.1038/srep07383
To quote:

Our findings suggest that highly toxic wavelengths of visible light
are species-specific in insects and that shorter wavelengths are not
always more toxic. For some animals, such as insects, blue light is
more harmful than UV light.

Granted they irradiated the insects for hours or days, but the point is that visible light can be very harmful for some insects (including, specifically, Culex pipiens molestus!)

Yes, the Sun is a very powerful source of light, but it's not concentrated. Like @canadianer said, the intensity is very high if all the light from a torch is forced onto a small space... and this is precisely what's happening (I specifically mentioned "white ceiling" because it causes all the light to bounce back on the insect!) When the torch is nearing the ceiling, the reflection gets so bright that it's very painful to stare at it (more painful than staring at the Sun)!

Ability or inability of a torch to burn skin has nothing to do with lethality. There might be many mechanisms behind this effect; I suspect their ganglion is overloaded by the intense light and it switches itself off!

And, finally, "common-sense logical answers" such as "insects fly all day in the sunlight and don't die" is not something I'd expect on SE. If you think that my conjecture is wrong, I'm fine with that, provided that you quote articles and list their DOI's.

I beg to differ @jakebeal. There're a lot of space between flashlight's emitter and the ceiling, the insects are buzzing and moving around. Moreover, they're not crushed, they look intact (even medium-sized flies, mosquitoes are much smaller but still they dropped dead!)

That laser weapon is physically damaging the insects @jakebeal; agreed, a handheld torch won't be powerful enough to do that. However, I was thinking that intense light simply overloads their CNS, it short-circuits and they drop dead! :)

@Alexander: Yet it's common to see mosquitos & flies flying around in broad daylight, which is much more intense than any flashlight I've heard of.

@jamesqf If you have a 1000 lumen flashlight held close enough to a surface such that it illuminates 0.001 square metres (about 3cm x 3 cm), the light intensity on that surface would be 1,000,000 lux. According to Wikipedia, that is about 10 times more intense than “bright sunlight”.

include the model of the flashlight and the distance and this can be answered to your satisfaction, otherwise all that is possible is just anecdotal conjecture.

Did you verify your insects were dead rather than momentarily blinded or confused into immobility?  This could make for a popular video, provided the flashlight doesn't kill the camera. :)

Without knowing the specifications of your flashlight (other than "very powerful") it is difficult to answer this question, however there is one common feature of all forms of lighting; heat output.
Most of the output of flashlights (including LED) is in the form of heat - more than 50% in the case of LEDs, greater amounts for incandescent bulbs (>95% IIRC). I've found a little information on this without diving into the literature too much.

Although LEDs are cool to the touch, within the devices themselves, there is plenty of unwanted heat. This heat comes from the inefficiency of the semiconductors that generate the light. The radiant efficiency (total optical output power divided by total electrical input power) of LEDs is typically between 5 and 40%, meaning that 60-95% of the input power is lost as heat.

I don't know how much of the heat is radiated out the front of the flashlight as this will depend on the flashlight design.
It's likely that you just got the insects heated to a point where they died from heat.

Why the downvote - it is the correct answer, nothing to do with biology or "white" light output.

For what it’s worth, I have an LED flashlight the manufacturer claims is 1800 lumens and I can’t hold it next to my skin for more than a few seconds. Shining it at my skin from an inch or two away and it still becomes too hot in 5-10 seconds.

@canadianer - I've had a minor burn (sore and red) on my leg from the LED flash/flashlight on my phone camera that was on in my pocket and I was unaware - that's going through the pocket material, so I am not surprised that an 1800 lumen one is hot to the touch.

I didn't downvote @bob1 :) Hmmm maybe... meaning that the insects are not heated directly from the contact with the bulb (because I can easily touch and hold it), but maybe the light heats them up instead... My torch is 1600 or 1800 lumens but it's a diving torch with a very effective radiator, so it gets hot only after a couple of minutes. When I switch it on, it's cold to the touch.

downvoted because it is not "likely" at all, you would at least have to show the heat output is at a dangerous level which is difficult without knowing the intensity, distance, and exposure time.

I think what is missing here is the proper experimental procedure, where all the effects, potentially harmful to insects (heat, physical pressure, etc.) are controlled. Think about making a movie of your experiment, doing it in different places and on different days. As now the discussion is meaningless.
Designing such an experiment could be an interesting question in itself, although I am not sure whether it is more appropriate for this forum or for Cross-Validated (statistics).
Note also that visible means a particular frequency range - the one perceived by our eyes. The frequency by itself is not harmful - what may cause harm is heating due to the energy absorbed with the light (microwave, infrared and ultraviolet radiation are known to heat not less than the one in visible range). Energy transmission with incoherent light (as opposed to the coherent laser light) is very inefficient, so it is unlikely to be the cause... yet, scientifically one cannot make any conclusions on the basis of the evidence presented so far.

The first experiment could be to simply use the torch while switched off. Who knows, maybe it's simpy the intense tries of the insects to flee - they might literally be "hitting their heads" on the glass and stunning themselves due to impact-

actually all you need to do is show insects survive higher intensity light just fine, which you can do just by knowing the flashlight involved.

@AnoE Is an insect actually capable of stunning itself?

@John It depends on the question one poses in the experiment: if one wants to test whether it is the  that flashlight kills the insects, than this could be the first step (because, if it can kill the insects, it will still remain to prove that this is the case). If one wants to clarify why they die (and whether they die), it might exlude the flashlight hypothesis.

@RogerVadim, that's the question. The scale of "physical impact" has two extremes: 1) no impact, and 2) the insect hitting my car's windshield at 200km/h. We know what the latter does to the insect. So somewhere between zero and 200 km/h there is a point where the insect will be almost, but not quite dead.  I have no idea whether insects can reach this kind of kinetic energy by hectically buzzing around in an extremely constrained space, but generally nothing surprises me anymore when hearing this kind of stuff... this would explain OP's question if it could be demonstrated in experiment.

@AnoE Another possibility is that the insects in question are not dead, but merely disoriented/paralized.

No, if the flashlight cannot harm your skin its is not killing an insect. In the absolute best case scenario you dazzled the insect temporarily in the same way intense light can dazzle you. lasers have been used to kill insects and they are quite intense, low intensity lasers far more powerful than your flashlight are used to identify insects and they do not kill them. Can you kill insects with intense visible light, absolutely, but your flashlight is nowhere near powerful enough.

"if the flashlight cannot burn your skin its is not killing an insect". Is it necessarily the case that insects require more energy to be killed that human skin does to be burned? I think some kind of logic or reference would be good.

@user438383 both are made of the same material, mostly water, and sunlight can burn human skin yet insects survive it just fine. lasers which can kill insects are quite painful on skin. keep in mind sunlight can burn human skin but will not kill most insects.

Like I said in comments, I don't believe either heat or the light itself is physically damaging the insects or their chitin carapace. In my opinion their brains are overloaded by the light...

@Alexander how do you think light can overload a brain? you have dazzling which I already mentioned, is temporary, and is not lethal. If your propose some other effect you would have to demonstrate such an effect even exists. And again sunlight is brighter and does not "overload" anything.

Hey @John, well it's called Sensory Overload in humans... loud sound, bright light or other stimuli can cause serious adverse reactions, even though they don't cause physical harm. I guess nobody died from it, but insects are very different from us...

@John, like some people said, the sunlight is diffused but torchlight is very concentrated and reflected from the white ceiling. Anyway, I'm not trying to prove anything, I just made an observation and asked a question and am addressing "obvious" answers such as "insects are crushed by the torch" :)

you said it yourself sensory overload (dazzling) does not cause physical harm. Also I don't think you understand how light intensity works as how diffuse the light is means nothing if the light intensity is higher if anything that means the insect is receiving even more energy from sunlight.


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Epigenetic marks
I was lately researching epigenetics and the current research on age reversal. Because epigenetic processes are influenced by the environment and can vary over time, I wanted to know if all epigenetic marks and mechanisms, including those that play a key part in cell differentiation, are reversible?
I am extremely keen to learn about this and any assistance would be greatly appreciated.

Welcome to SE Biology! When you say "reversible", what do you mean exactly? All chemical processes are theoretically reversible. It would help to know more about what specifically you have read, and what does not answer your question from e.g. a google search or wikipedia page. For example, I find multiple resources on this topic by google (https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3411986/, https://www.ncbi.nlm.nih.gov/pmc/articles/PMC5137275/). Is there something specifically that you don't understand? Otherwise this question will be difficult to answer as it is quite broad.

I second the comments of @MaximilianPress but would ask you whether you know what “epigenetic tags” are in chemical terms. Because if you do not, you need to find out before you are in a position to ask here about their specific reversal because it is unlikely that you will understand an answer. Do you?

@David Sir, yes I studied them  and I understand them

@Maximillian Press Sir, I was curious if the epigenetic processes that drive cell differentiation and aging may be reversed. Can we undo all of the aging-causing epigenetic mechanisms?

@AIA that is good information to have in the question, but it is still very generic. There are many kinds of changes that people call epigenetic that play roles in development/differentiation, and yes they are reversible in some sense at the biochemical level. However, this does not necessarily have any relevance to aging. It would be helpful to know more specifics about what you have read and what specific questions remain unanswered. This is too broad to answer as is.

@Maximilian Press Sorry sir, I am not delivering clear information. Basically, I am making a video for a competition on the topic of slowing down and reversing aging with epigenetics. I read David Sinclair and other scientists' researches about it. So I just want to clarify that if we are reversing aging, so it mean that we are changing our epigenome and reversing some epigenetic marks? so I want to clarify is this right. Sir, if you have references to any research articles that would help me in getting great insight into this topic, i will be thankful if you can share them

To some extent, there are people who believe that we can reverse aging by reversing epigenetic marks. I don't know that I believe this, but there is certainly an academic literature on it. Notably, some of the same epigenetic marks associated with aging are also associated with tumor suppression, so reversing marks is not necessarily always a great idea. Here is a link to an article, that in turn links to a study. I wouldn't necessarily trust it, but it should lead you to more information: https://www.nature.com/articles/d41586-019-02638-w


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What species of viper is this?
I think that the snake in the pictures here below is a viper. I found it on a hiking trail on a mountain in Bulgaria. Height was about 2000 meters and the weather wasn't warm as you might expect in a late Spring days, but it seemed quite active.
If it really is a viper what species of viper? Otherwise what could it be?

This is most likely the common European adder (Vipera beris). A list of the reptiles of Bulgaria lists it as occurring in the mountains of Bulgaria up to 2700 metres.
There are a couple of other contenders in the list of vipers, but two of them are incredibly rare (not found since the 1930's) and the last is the horned viper (Vipera ammodytes), which has distinctive horns on the tip of its nose.

Thank you. The description of the common adder is a good match and it had no horns, although it was quite young I think the second option can be discarded.


text:
Isolating colonies in a pure culture
I want to conduct antimicrobial susceptibility tests for which I've to culture bacteria first. Since I've bought KWIK-STIKs, (KWIK-STIK contains a single lyophilized microorganism which have to be rehydrated before culturing them on agar plates) I referred to this video for instructions.
In the video, the bacteria are rehydrated and streaked on the agar plates to obtain individual colonies. I know one should isolate colonies in order to obtain a pure culture. But why should you isolate colonies if the culture was already pure?
Here, I'm assuming that companies sell bacteria which are already isolated and sampled, therefore are pure (that's why I wonder why isolating colonies is required), but I'm not a professional in the microbiology field (I'm a high school student) so any basic clarification would be welcome.

Your question is not stupid, but it is not clearly expressed. Please rewrite it. Start by explaining clearly what you are doing. Colonies of what? What kind of culture. What is your aim? You refer to a video, but nobody here knows anything about it, and giving us a link is no good because questions here have to be self contained. So you need to carefully report what the video advises, quoting the advice verbatim. That’s how science  works.

@David Ok, I've rewritten it...is it clear now?

Yes that's better. Someone with the appropriate background will have something to go on now.

The answer is that it allows you to check viability of the stock primarily, and thereby also have a population that you know is growing. Bulk cultures from liquid will always contain a proportion of non-viable organisms as there are no easy ways to separate these from the viable.
It also allows you to check to see that the preparation hasn't been accidentally contaminated (or tubes swapped etc.) by another morphologically different colony former - having the right type of organism is very important experimentally. If you wished to check the morphology of the organism by either morphological (i.e. microscope + stains) or biochemical tests, both of these need to start with a pure culture. It is hard to get enough bacteria for most of these tests unless you pick them from colonies - liquid culture just isn't concentrated enough, and can be heterogenous if a mixed inoculum is (accidentally or otherwise) added.
Another reason is that there is some heterogeneity in the culture anyway, no matter what you do to characterize it after growth. Isolating from single colonies, which should be derived from single colony forming units, allows you some control over this population for your bulk preparation, whether this be growth under certain conditions (e.g. antibiotic resistance) or growth rate.
It for instance may allow you to select a large colony, which might indicate a slightly higher growth rate for that founder. If you have something that is grown on selective media, then it may be that a portion of a bulk culture has lost the characteristic for growth on this medium, which means that when you inoculate from the bulk, what you put in might not be what you expected. This can lead to deviations in expected generation time calculations and similar physiological or phenotypic measures.


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I had an encounter with a cougar (mountain lion). What does the recurrence time look like?
I had an encounter with a cougar recently. I was walking down a muddy old forest service road when approximately 15-20 meters behind me a deer burst out of the brush with a cougar chasing it close behind. The cougar gave up the chase on the deer but watched it jump away (instead of turning around to see what I was doing). Meanwhile I am backing away around the corner to get out of the situation. I didn't see either the cougar or the deer again.
What I'm curious about that a biology perspective might be able to answer is how frequently cougars return to a hunting area. My qualitative understanding is that they have very large ranges, so under a quite few stochastic process models I would expect them to return to the same hectare relatively infrequently. But I don't have a quantitative understanding of this.
Has someone worked out some empirical distribution (conditional or marginal) on the return-time of cougars to a given hunting location?
Update
A cougar was spotted by someone else approximately 15 kilometres away within 2 weeks of my encounter.

This observation is a Casual observation on iNaturalist: ID 110726469

I'm guessing the variance on something like this is too great to be useful, beyond what you could roughly estimate by range. But perhaps someone will be able to answer anyways.

Yes, the variance could be very large. I also find that ecological models are not adequately validated, and rely too heavily on either linear preasymptotic AIC or asymptotic AIC to be realistic. But, I will take what I can get.

An obvious (at least to me) problem with doing such an analysis is that it assumes that all locations within the range are equally favorable.  But deer (a preferred prey of the cougar) have their preferred trails & feeding areas, so a successful cougar is likely to learn to hunt where the prey is, and not where it's not likely to be.

After a bit of googling: home range of a cougar is between 30 and 1000 km². If we assumed that it is a circle, the bigger number corresponds to a diamater of just above 60 kilometers, which is a two-three days walk for a human in a forest. So it is actually not that big.

@jamesqf In the case of a marginal distribution, yes. You can have conditional probability distributions that are conditioned on things like sources of water, trails, shelter, etc. Obtaining such detailed data is considerably more work.

@RogerVadim Thanks for looking that up, and for the back-of-the-envelope calculation. It raises interesting questions about the geometric properties of the home ranges of animals.

It did not answer my question, but this article on Fourier analysis of animal location is quite interesting.

this file gives the raw step-lengths and turning angles of a bunch of Florida panthers (from this paper). You could start there ...

@BenBolker Thanks! Considering another big cat is an interesting heuristic for representativeness.


text:
Were the Francevillian biota real?
The announced discovery of macrofossils suggestive of large multicellular organisms 2.1 Ga ago is now over ten years old.
Is there by now a scientific consensus whether these constitute real fossils of an abortive early evolution of complex life, or real fossils ancestral to extant life, or simply pseudofossils just looking like complex life?

Interesting question, you might be better off posting it to the Earth Science SE site for this one. All the papers discussing these fossils were all from the same authors, so I can't say one-way or the other.

Yes they are real.
it is a a collection fossils that are different enough from modern forms it is difficult to say exactly what they are but they definitely exist. Occasionally someone proposes a specific individual fossil might be Pseudofossils but so far they have held up to a great deal of scrutiny. Some are too regular to even be disputed. much like the Ediacaran they are just forms very different from anything we are used too.
https://journals.plos.org/plosone/article?id=10.1371/journal.pone.0099438

I'm also a believer in them, but I'm interested to find out on what basis you found that the scientific community put them under a lot of scrutiny? All I could find were papers published by the same group, admittedly largely in well respected journals, but nothing showing any form of debate - which may in itself be telling of acceptance.

don't expect to find a lot of authors of papers from different places for fossils. Funding, physical access, ad the lack of paleontologists  tend to mean only those very interested publish papers especially on invertebrates.

Thanks. That makes sense, I had wondered if the situation were something like that - I don't imagine a whole bunch of people outside of geographically close countries get to Gabon in general.