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Does natural selection still increase biological complexity?
I recently read The Selfish Gene by Richard Dawkins, which I found very interesting. In one of the last chapters, he gives multiple possible explanations to the question "Why did natural selection give rise to complicated lifeforms?".
Since complex (say: multi celled) life exists, that means natural selection at some point did increase complexity, as we went from the primal soup to multi celled organisms (with many steps in between). My question is whether this trend of increasing complexity is still going on? Is natural selection a never-ending arms race, tending towards increasingly complicated genomes? Or did natural selection reach some plateau, local optimum, of biological complexity, after which the only evolution taking place is "on the same level".
I have thought a bit about a metric for complexity, but since I have no background in biology I am not at all certain. At first thought, the length of the genome sounds like a promising candidate, but the phenomenon of freeloading, not-functioning DNA (conveniently described in The Selfish Gene too) undermines this. I hope it is clear what I mean by "complexity".
Thanks for answers in advance, it has been fun speculating.

Hi Lourens & welcome to Bio.SE! This is an interesting topic. To make your question more answerable, try to focus in on a specific question. If you could start with your definition of "complex-life" and ask a question on what you don't understand about the definition. For example, your current definition is simply how "multicellular" an organism is. Is a blue whale more complex than a human?

On a side note, the Selfish gene is >40 years old. We understand far more about genes now than we did then. For example, even the definition of "non-functioning" DNA is problematic; wikipedia describes this evidence of functional "junk" DNA.

@James yes specifically the definition of complexity is difficult, as I mentioned too. But we can agree that a human is more complex, then, say a microbe. So at some point, natural selection gave rise to more complexity. What my question boils down to is whether this is still happening.

Answering that last question depends intimately on a definition of complexity. I'm not quite sure where people get so interested in complexity and evolution, but questions similar to this have been asked numerous times here and they have all failed (as far as I recall) to give any operational definition of complexity that can help answer the question.

People seem to forget that bacteria and archaea have been evolving for just as long and they largely remain unicellular.

Possible duplicate  Is variation a result of Evolution? or https://biology.stackexchange.com/questions/42050/if-evolution-is-not-about-increased-complexity-why-does-so-much-complexity-evol

Sure, a human seems to be more complex than E.coli. We would start answering this question by trying to measure & quantify the complexity of these organisms. Is the complexity the number of cells, something anatomical like the number of appendages, environmental adaptability, variety of hats, number of coding genes, or sequence complexity? Only then can we discuss if that specific feature appears to be rising, falling, or plateauing in the biosphere. Without these specific exact definitions, this question cannot be answered completely.

There are multiple definitions of complexity, ranging from algorithmic complexity to computational complexity.  In biology, I'd say that algorithmic complexity (Kolmogorov complexity) is most relevant.  Unfortunately, we don't yet know enough about genetic circuits and all the other things that go on at the subcellular level even to make plausible estimates of an organism's complexity.
If the complexity of a human being were analyzed all the way from physiology down to the subcellular & molecular level, I think it would be apparent that most of the complexity is present at the subcellular & molecular level.  In fact, essentially all of the complexity of a multicellular organism is encoded at the subcellular level, so it's appropriate to say that's where the complexity resides.
It is not entirely clear that a human being, whose genome contains ~3 billion base pairs, is more complex than a protozoan Amoeba Proteus whose genome is ~34-43 billion base pairs.
All of the processes that have been active in evolution since the beginning are still active, and changes in complexity - regardless of the definition of complexity - are still going on.
The closest things there is to a "plateau" is a relatively static distribution of genotypes that can last only as long as the adaptive landscape is unchanged.  However, the adaptive landscape is always changing because of climate changes, plate tectonics, invasive species, disease, intra-species iteractions, and even because of chance fluctuations in the distribution of genotypes.

thank you for your answer. As @James also pointed out in a comment, my question fails by the definition of complexity. Not only that, the abstract idea of complexity I had in mind seems flawed after reading your answer, since I was clearly biased towards the human scale. Your fact about the amoeba dubia's genome seems to be outdated, although your point still stands. Correcting for the measurement error gives the indirect estimate of 67 billion base pairs. https://bionumbers.hms.harvard.edu/bionumber.aspx?id=104470&ver=6


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What is the linear force in a bicep compared to the force at the hand?
So I know that like most joints, the bicep applies a force near to the pivot (the elbow) and creates a rotational torque in the elbow, this moving the lever, your forearm and hand. Due to the biceps close proximity to the pivot it at a considerable mechanical disadvantage compared to a downward force on the hand meaning linearly, muscle are a lot stronger than we think. Assume somehow you could take out a bicep, for example mine that has a max curl of 25kg dumbells, and hung it from an anchor point, removing the pivot. What would be the maximum linear force of that muscle? E.g. would it be 4x stronger than the torque in the elbow that's applied at the hand and therfore lift 100kg?
Sorry this question is a bit confusing. I really didn't know how to describe this hypothetical scenario. I hope you understand what I am trying to ask.

Are you essentially asking for the summed force of all myosin-actin cross bridges of an average bicep outside the context of the body?

Also, you must recognize that the strength of a person's bicep can vary greatly between different people -- wouldn't you expect such variability to also exist in terms of your question's scenario?

Lastly, given my two above comments, might I suggest you rewrite/rethink your question slightly differently? I suggest focusing on the pulling force of a single actin-myosin cross bridge. You could simply multiply whatever value that is upward based on some average number of cross bridges per sarcomere/myofilament/myofibril/fiber/fascicle

My answer is in the context of my comments posted under the main question. I tried scaling up forces throughout the muscle. I probably flubbed one or more things, so check my math and suggest fixes!!
The short answer:
The max linear force of an average adult male bicep is ~1600 Newtons.
According to Lumen Learning, the force ratio between bicep and hand when lifting 4kg is ~7.4. (note that my example would result in a ratio closer to 9). Regardless, using their free body diagrams and force equations, my estimate of ~1600N is the equivalent of lifting ("curling") ~16kg (35 lbs) held in the hand. That seems reasonable, so, my math is likely in the right ball park!
The longer explanation:

VanBuren et al (1995) estimates ~0.2 pico-Newtons per cross-bridge.

In a direct comparison between smooth and skeletal muscle myosin, the average force per cross-bridge was 0.8 and 0.2 pN, respectively. Surprisingly, smooth muscle myosin generates approximately 4 times greater average force per cross-bridge head than skeletal muscle myosin.

The multiplication*:

Note: Muscle fiber anatomy has been detailed quite robustly: AL-Khayat 2013, Wang et al 2018, Mijailovich et al. 2019

Ojima 2019 claims each thick filament is formed of approximately 300 myosin heads. (though, I've seen less reputable sources [e.g., here] suggest as high as 500 without further support).

Ojima 2019 also claim each thick filament is ~15nm in diameter and 1.6µm long. AL-Khayat 2013 agrees on length, but suggests this could be as large as 30 nm in diameter. Szikora et al 2022 claims the length is consistent/conserved around 1.65μm across vertebrates.

We'll stick with the minimum suggestions: 15 nm x 1.6µm.

Also, note that AL-Khayat 2013 suggests actin filaments are ~10nm in diameter.

Higuchi et al. 1991 suggest myosin heads are 19nm long,while AL-Khayat 2013 suggests 15nm each.

Gerling & Brown 2013 found sarcomere lengths (at least in human latissimus dorsi) to have an average length of 2.7μm, while Dasbiswas et al 2018 suggests "The sarcomeres in relaxed human skeletal muscles are between 3.0 and 3.4 µm long, depending on the muscle sub-type." So, let's assume 3µm.

Previously, the force produced by a skeletal muscle myofibril was assumed to depend on cross-sectional area alone and not on the number of active sarcomeres (because of their arrangement in series). However, Mendoza & Rassier's (2020) results suggest that myofibril force is affected by intersarcomere dynamics and the number of active sarcomeres in series. As such, the total number of sarcomeres per volume of muscle is likely important.

AL-Khayat 2013 suggest vertebrate skeletal myofibrils are typically 1–2 μm diameter and 2–50 mm long.

Regarding Diameter:

Smith & Plowman 2007 note: "in regions where the thick and thin filaments overlap, each thick filament is surrounded by six thin filaments and each thin filament is surrounded by three thick filaments." If each thick filament is 15nm diameter, each thin filament is ~10nm in diameter, and each myosin head (i.e. space between filaments) is 15 nm, it would suggest each myosin and actin complex would be approximately 65nm in diameter. That would be a maximum of 15-30 actin-myosin complexes per diameter of myofibril. So let's stick with a conservative 15.

Regarding length:

If sarcomeres are an average of 3µm in length, 2–50 mm long myofibrils suggest an average of 667 - 16,667 sarcomeres per myofibril. The average is ~8600 sarcomeres per length.

Putting length and diameter together, this would suggest that each myofibril might contain 15 x 8600, or ~130,000 thick filaments on average.

130,000 x 300 = 39 million myosin heads per myofibril.

Chen et al 2017 state: "the number of myofibrils ranges from 50 per muscle fiber in the muscles of a fetus to approximately 2000 per fiber in the muscles of an untrained adult."

This would suggest 39 million x 2000, or 78 billion myosin heads per muscle fiber.

There seems to be a couple hundred thousand fibers in human biceps:

Klein et al 2003: "The estimated number of muscle fibers in the biceps ranged from 201,360–312,708 in the young men and 149,301–291,501 in the old men"

MacDougall et al 1984: "There was a wide interindividual range for fiber numbers in biceps (172,085–418,884), but despite large differences in muscle size both bodybuilder groups possessed the same number of muscle fibers as the group of untrained controls"

The average is around 280,000 fibers per male bicep.

280,000 fibers x 178 billion myosin heads per muscle fiber = 2.2 quadrillion (2.2x1016) myosin heads in a biceps.

This equates to 2x10-13 N per cross-bridge x 2.2x1016 cross-bridges possible, or 4400 Newtons.

But, Mettikolla et al 2010 suggests only about 1/3 of cross-bridges are active at any time during isometric contraction:

These results imply that cross-bridges interact with actin in bursts and suggest that during active period, on average 36% of cross-bridges are involved in force generation

This suggests we should divide our above outcome, 4400 N, by 36%:
As a result, my math suggests ~1584 N per bicep

*MAJOR CAVEAT: Moo et al 2016 suggests that my above approach is faulty:

Sarcomere lengths have been a crucial outcome measure for understanding and explaining basic muscle properties and muscle function. Sarcomere lengths for a given muscle are typically measured at a single spot, often in the mid-belly of the muscle, and at a given muscle length. It is then assumed implicitly that the sarcomere length measured at this single spot represents the sarcomere lengths at other locations within the muscle, and force-length, force-velocity, and power-velocity properties of muscles are often implied based on these single sarcomere length measurements. Although, intuitively appealing, this assumption is yet to be supported by systematic evidence

We showed that sarcomere lengths varied substantially within small regions of the muscle and also for different sites across the entire [muscle]. Also, sarcomere elongations with muscle lengthening were non-uniform across the muscle, with the highest sarcomere stretches occurring near the myotendinous junction. We conclude that muscle mechanics derived from sarcomere length measured from a small region of a muscle may not well-represent the sarcomere length and associated functional properties of the entire muscle

Is my answer reasonable?
I think so based on the link shared by the OP from Lumen Learning, which calculated a linear force of 470N to lift 4kg. Using their force equation, 1600N would be required to lift 16kg,or ~35lbs.that's about in line with what I see a typical healthy/athletic male pulling off while dumbbell curling at the gym!
Biology and physics agree!!

This link I found answers my intended question fairly well and simply.
https://courses.lumenlearning.com/suny-physics/chapter/9-6-forces-and-torques-in-muscles-and-joints/

source: Stack Exchange

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How lethal is high dose UV?
I watched a Neil DeGrasse Tyson video about teraforming Mars where he goes over the topic of UV radiation and how our ozone layer protects us. He makes a statement that the UV on earth is actually only a small percentage, about 3%, and furthermore only 5% of that being the more harmful UV-B which causes sunburn and skin cancer.
My question is, what would happen to a human if they were to receive the full dosage of UV-B radiation from the sun, let us assume 500 times higher than that on earth, with no protection? Would it be fatal? And if so how fast would it kill you?

The answer is easier than you think. Have you ever had a sunburn so bad that it causes blisters? (I have, as a teen falling asleep on the beach.) Those sunburns are called second degree sunburns. With that amount of exposure, the least that would happen is a 2° sunburn over most of your body (some parts don't suffer this), which would lead to hypovolemic shock. Without treatment (which includes fluid resuscitation), you would probably die within 24 hours. If, by chance you survived the first 24 hours, the blisters that break open (and they would) would get infected because your immune system would be compromised by the damaged capillaries of the skin (that's one cause of the blisters in the first place.) Your lifespan would be a few days if you didn't die within the first 24 hours.
This is the best case scenario.
This is why burn units exist. Second degree burns over a certain percentage of your body are lethal.

You wouldn't get something like radiation sickness but something can be so unhealthy that can then lead to increased mortality. Whether or not something kills you is the lowest bar that can be set for whether something is healthy or not.
I couldn't actually find high wavelength resolution graphs for the solar irradiance at sea level since the values are vanishingly small on normal graphs.
However, if you take the the UV irradiance at outside of the atmosphere at 250nm to be 0.25W/m^2/nm and use that as a baseline for the UVB range of 280-315nm, then the irradiance 8.75W/m^2. Five hundred times that is equivalent to a 4.375mW UV laser producing a 1mm x 1mm square pot. The first thing you would notice is that you would go blind.

https://www.e-education.psu.edu/meteo300/node/683

Not a bad answer. For actual conditions on Mars: Mars is about 1.5 AU, so by inverse squared law = 1/1.5^2 = 0.44 so irradiance is 0.44 of Earth.

It depends on the wavelength. This paper1 provides the numbers:

This study was conducted to determine the toxicological effects of different wavelengths (250, 270, 290, and 310 nm) and doses of UV radiation on cell viability, DNA structure, and DNA damage repair mechanisms in a PC12 cell system. For this, we evaluated cell viability and CPD formation. Cell survival rate was markedly decreased 24 h after UV irradiation in a dose-dependent manner at all wavelengths (except at 310 nm). Cell viability increased with increasing wavelength in the following order: 250 < 270 < 290 < 310 nm. UV radiation at 250 nm showed the highest cell killing ability, with a median lethal dose (LD50) of 120 mJ/cm2. The LD50 gradually increased with increase in wavelength. Among the 4 wavelengths tested, the highest LD50 (6000 mJ/cm2) was obtained for 310 nm. CPD formation decreased substantially with increasing wavelength. Among the 4 wavelengths, the proportion of CPD formation was highest at 250 nm and lowest at 310 nm. On the basis of LD50 values for each wavelength, PC12 cells irradiated with UV radiation of 290 nm showed maximum DNA repair ability, whereas those irradiated with the 310-nm radiation did not show any repair ability. Toxicity of UV radiation varied with wavelengths and exposure doses.

You can find more details on this topic in ref.2
References

Masuma R, Kashima S, Kurasaki M, Okuno T. Effects of UV wavelength on cell damages caused by UV irradiation in PC12 cells. J Photochem Photobiol B. 2013 Aug 5;125:202-8. doi: 10.1016/j.jphotobiol.2013.06.003
Lawrence, K.P., Douki, T., Sarkany, R.P.E. et al. The UV/Visible Radiation Boundary Region (385–405 nm) Damages Skin Cells and Induces “dark” Cyclobutane Pyrimidine Dimers in Human Skin in vivo. Sci Rep 8, 12722 (2018). DOI: 10.1038/s41598-018-30738-6

I'm not sure you're addressing the OP's question directly, which involves damage to and even death in humans exposed to UV B radiation (280–315 nm), not skin cells in vitro or in vivo.

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Why does activity cause hypoglycaemia in diabetics?
(I'm from the UK so I'll be using mmol/L as units and taking <4.0 as hypoglycaemia.)
I remember being told that I should consume a small amount of sugar or reduce insulin dose sightly to increase blood sugar before any considerable physical activity. Only over the last year or 2 have I had to use this strategy regularly since starting at a gym. I remember being told that it was recommended to be >8.0 before exercise but in my own experience >10.0 is required to avoid hypoglycaemia after exercise. I'm simply asking why this is necessary. If my body does not produce insulin, how does the introduction of physical activity cause my blood sugar to drop? Are my cells absorbing the glucose without insulin? If not where is the glucose going?

While doing physical activity you are using your skeletal muscles. When the skeletal muscles are active i.e. contracting and relaxing during exercise, especially moderate or severe exercise, they (skeletal muscle cells) can take up glucose present in the blood, in the absence of insulin which will be used as a source of energy for the contractile process.
Thus there could be decrease in Blood sugar levels(BSL), i.e. hypoglycemia especially if you are taking insulin or any other tablets to maintain your BSL as you a diabetic. The dosage of these medicines/tablets is determined by assuming that you will be doing routine day to day activities.Any increase in quantum of physical activity such as exercise will result in hypoglycemia i.e. decreased blood sugar level during or after moderate/severe exercise.

Ref. Guyton, Textbook of Physiology.

Glucose is the primary fuel of cells. If you're diabetic and under medication, the doses you receive are calibrated for normal physical activity. If in case you engage in any unusually excessive physical activity, your blood glucose will fall due to two reasons:

The unusual physical activity
The diabetic drugs you're on

This combo of glucose-level-reducing factors may cause your blood sugar to fall to dangerous levels.  In essence the drug dose is too high (relatively) for that particular time you're engaged in unusually strenous physical activity.
In a nondiabetic the glucose level is kept at normal levels by controlling the level of insulin in the body + nondiabetic people's cells are responsive to insulin.
Last I checked, diabetic drugs are intentionally called hypoglycemic agents so that prescribers and patients are aware of the danger.


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Does cooking kill all bacteria?
So I know that the general rules is that pathogen prone foods such as chicken shouldn't be consumed after 2 hours at room temperature. Cooking meat such as chicken is necesary as it kills the potentially harmful bacteria in th raw meat. What I do t understand is, if the cooking process kills the harmful bacteria, why would it still be not recommended to cook and eat chicken left at room temperature for longer than 2 hours? Would the process of cooking not still be able to kill the increased amount of bacteria?

There are (at least) two questions here, do you want to pick one and ask the other(s) in a separate thread. Remember, show your research when posting.

Heat-stable toxins.

It would kill the bacteria, but it wouldn't break down the toxic substances that the bacteria produced while they were digesting the chicken.
According to the WHO, the chicken must be maintained at 85⁰C for at least 5 minutes, so it is possible to cook it, but there's no real way to know for sure that all the toxins have been destroyed without reaching temperatures that outright destroys the chicken.
Since there's no way to perfectly know whether the chicken has been cooked long enough to destroy the toxins, without turning it to ash, it's better to avoid the risk in the first place, since a small amount remaining could lead to unpleasantness, and the temperatures would overcook the chicken (Poultry is considered cooked when its interior temperature reaches 74⁰C).


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optimal spatial arrangement (distribution) of plants
Microalgae are organisms that can grow in aquatic environments and use light and carbon dioxide (CO2) to create biomass. Research showed that Microalgae's biomass can be used as a bio-fertilizer.
My mission is: to find the optimal field patterns . I mean the optimal spatial arrangement (distribution) of plants in a given area. To elaborate more, I need to consider if I plant a given land in raw (strip) pattern or rectangular pattern or maybe hexagonal or circular pattern, can this boost the crop yield or optimize the bio-fertilizer or the water management.
Note: I'm not a biologist, Just a data science student. I'm open to any biology/math resources and any other inspirations or approaches.

Is your mission to find the optimal pattern or for somebody else to find it for you? Also, can you please [edit] your post to clarify your specific question? Thank you.

yes it is my mission to find the optimal pattern. and my question is exactly : how to find this pattern for a given plant. thank you for your response@theforestecologist

Do we assume all "leaves" are on a flat surface? I.e., no height/leaves above other leaves.

we can assume that plants are planted at the same time so we can expect the same growth rate.  So yes we can assume that no height/leaves above other leaves. Sorry for my late response. @BigMistake

Perhaps the only way to find this out is to try different things and see what works.


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How to read cladograms/phylograms?
I understand the difference between cladograms and phylograms; the latter can show the evolutionary linkage over time. However, I am having trouble finding out which two organisms are most closely related.

In the image above for example:
Referring to the cladogram, would these following statements be correct
-Bacteria 1 is least related to Eukaryote 4 (the furthest branch is always the least related as there common node is furthest away, correct?)
-The most related organisms are Bacteria 2 and 3, and Eukaryote 2 and 3
Referring to the phylogram
-Is Eukaryote 2 the least related to Bacteria 1 as its branch is the furthest away? Or is it Eukaryote 4 as it is the furthest away (bottom of list)?
-Is Eukaryote 1 most closely related to Eukaryote 3?

I hate to be pedantic, but it should be bacterium (singular) rather than bacteria (plural). I know few people know any Latin any more (including the originator of the diagram), but just so you know next time. After all science is about precision.

@David I didn't make the image, but doesn't it work either way since there are many organisms of each species...

Yes, I know you didn't make the image, and appreciate your difficulty. (I would have modified it myself, but I do that sort of thing.) However I think you would always use the singular (as in eukaryote). For example if you substituted 'animal' or 'microorganism' where the plurals are English,  'animals 1' or 'microorganisms 2' would strike one as odd. You certainly wouldn't want to mix them.

-Bacteria 1 is least related to Eukaryote 4 (the furthest branch is always the least related as there common node is furthest away, correct?)

In terms of relative relationships, eukaryote 4's least-related relative is Bacteria 1. Bacteria 1's least-related relative is the entire rest of the tree

-The most related organisms are Bacteria 2 and 3, and Eukaryote 2 and 3

You cannot actually answer this question because a cladogram only shows relative relationships. To know which species pair is the most related (has the least total distance between them), you would need to know the branch lengths.

-Is Eukaryote 2 the least related to Bacteria 1 as its branch is the furthest away?

To determine relationships, you want to look at the total branch length between two species. Therefore, this is correct

Or is it Eukaryote 4 as it is the furthest away (bottom of list)?

Avoid this way of thinking. A phylogeny is not a 'list'. Pay no attention to the order the species labels appear: you only want to pay attention to how the species are connected. The same phylogenetic tree could be drawn differently, placing Eukaryote 3 at the bottom, for example.

-Is Eukaryote 1 most closely related to Eukaryote 3?

Eukaryote 1's closest relative is Eukaryote 4: the distance from Eukaryote 1 to Eukaryote 4 is the shortest out of all possible pairs with Eukaryote 1.

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Abdominal muscles during inspiration/inhalation
During inspiration I am aware that the diaphragm and the external intercostal muscles contract, thus increasing the volume of the thoracic cavity.
However, I read on my textbook that during inspiration:

the diaphragm contracts, and at the same time the external intercostal
  muscles and one set of abdominal muscles help to raise the rib cage.

What I do not understand is the movement of the abdominal muscles, as I thought that they help in ventilation during exhalation (contract during exhalation, but not during inhalation). Is my assumption incorrect?

In inspiration just the external intercostal muscles contract and in expiration the abdominal muscles contracts, and the external intercostal muscles relax thus the diaphragm relaxes and curves upwards. Furthermore, also in expiration, the internal intercostal muscles contract

Also note that during Eupnea, in inspiration only the diaphragm and external intercostal muscles are "required"

@JordiZambrino Okay, so what is written on the textbook is incorrect right? And during expiration the diaphragm relaxes? Thanks.

Not necessarily, its just that my textbook states something different. And for your second point: The Diaphragm relaxes  and curves upwards, the abdominal muscles contract, thus pushing the Diaphragm upwards, consequently the External intercostal muscles relax and the ribs fall, in the end, the Internal intercostal muscles contract, hence pulling the ribs downwards

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Blood antigens and immune response
In my textbook, the definition of an antigen is written as follows:
Antigen: A substance that the body recognises as foreign and that can evoke an immune response
The following image also confused me as it states that, for instance, a person with type A  blood has A-antigens

However, would this statement, and image, not be stating that the antigens found on the surface of our blood cells are foreign? Which would imply that our antibodies attack our own blood which is not what occurs?
Or are the substances found on the surface of blood cells (A or/and B antigens, no antigens) only recognised as antigens when they correspond with the antibodies in the body?
In other words, if I am blood type A, the pink molecules on the surface of my blood are not referred to as antigens as they are not foreign, but the green molecules on the surface of group B blood are referred to as antigens as they are foreign?

Which textbook are you referring to? Your body treats another person's antigen as foreign and it evokes an immune response. In this case, A antigen will evoke immune response in a person having B antigen or anti-A antibody as that anti-A antibody will treat A antigen as foreign. If you're satisfied by this, then I can post it as answer (though this is a homework question).

The problem is partly the definition of Antigen that you quote. It refers to “the body” leading you to assume it’s a specific body — yours. I'm sure there are better definitions, but just taking this one, I would modify to: “A substance that can be recognised as foreign and evoke an immune response in an organism.”
With this definition, which is actually how antigens are thought of, a particular antigen need not be antigenic in every organism to which it is presented. Those organisms or individuals for which the antigen appears as ‘self’ will not recognize it as foreign as they are ‘tolerant’ to it (e.g. by mechanisms which eliminate the cells capable of producing antibodies against the self antigen). But it is still referred to as an antigen.
So your understanding of the situation is correct, but that of the use of the terminology is not. Even if you belong to blood group A and therefore will not mount an immune response to the pink blobs in the picture when transfused with blood of group A, they are still referred to as antigens as they stimulate an immune response when transfused into a person of blood group B.
If you wished to talk about the fact that the A antigen on blood of type A does not provoke an immune response when transfused to persons with blood group A you could say “The A antigen on red cells is not immunogenic (or even antigenic) when transfused into persons of blood group A”.
Footnote on terminology
Scientific terminology allows us to describe things precisely. As our understanding increases, the terminology may need to evolve to allow us to make distinctions of which we were previously unaware. The word ‘antigen’ was coined in the late 19th century, before anything was known of the structure of antibodies, and was defined in terms of the immune response as that was what was observed. The definition evolved in some quarters to include the idea of binding to an antibody or T-cell (see e.g. Merriam-Webster definition).
However this extended definition raises the problem of the terminology to use if something binds to an antibody but does not provoke an immune response. Thus, the Glossary of the textbook Immunobiology (Janeway et al.) defines antigen as “Any molecule that can bind specifically to an antibody” and uses a different term, ‘immunogen’, for “Any molecule that can elicit an adaptive immune response on injection into a person or animal”. (Which explains my mention of ‘immunogenic’, as opposed to ‘antigenic’, above.)


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Which steps occur in each phase of Meiosis and Mitosis
MITOSIS

In mitosis am I correct in saying that the spindles attach to the centromeres at the end of prophase/prometaphase? (I read this in an answer to another question)

MEIOSIS

Does the nuclear envelope disintegrate during prophase (as occurs in mitosis), or during metaphase? (this is what it says on my textbook)
When do the spindles attach to the chromosomes during the first phase? (during prophase I, or anaphase I? On my textbook it says anaphase I)
And when do they attach to the sister chromatids in phase II? (on my textbook it says metaphase II).

Why would you doubt the answers you are reading in your textbook? It should be accurate, at least for a phenomena as well characterized as Mitosis and Meiosis.

Please limit yourself to one question per post.

Is there some reason that you doubt the textbook?

@rg255 I doubt the textbook because other reliable sources were giving contradicting information, and my teacher even mentioned that in some parts it was not specific/accurate enough.

Then you should give the sources for the both sides of the contradictory information

Mitosis:
1. Yes, the spindle fibers attach to centromeres at the prometaphase
http://www.biology.arizona.edu/cell_bio/tutorials/cell_cycle/cells3.html
Meiosis:
1. The nuclear membrane breaks down in Metaphase 1 and reforms during Telophase. It breaks down again in Prophase 2, reforming after that in Telophase 2 in the daughter cells.

The spindles attach to the centromere is Metaphase 1
Yes the spindles attach to the sister chromatids in Metaphase 2

Check out this animation, it might be pretty useful


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How do I classify cytotoxicity values, whether a sample is mildly, moderately, or highly cytotoxic?
I used LDH assay for cytotoxicity testing. I have a plant extract which I tested against HepG2 cancer cells. I did three trials, my results were 2%, 6%, and 8% cytotoxicity, respectively.
How do I classify these values?
I was wondering if these values are low/mild, moderate, or high cytotoxicity. Someone told me that anything higher than 10% is already highly cytotoxic; however, I cannot find any reference that stated this.

Welcome to Biology.SE. Please take the [tour] and then go through the [help] pages starting with [Ask] questions effectively on this site and [edit] your question accordingly. In particular, you should tell us: 1) what the LDH cytotoxicity assay is and what LDH stands for; and 2) if you are using a kit — from what company or if not, what protocol are you following (summarize and then provide references). Finally, please tell us what you have done to find the answer — as  it stands this seems under researched and possibly opinion-based, both of which are grounds for closure.

I hope the results are compared with the effects of the solvent alone on the cancer cells. For example, Ethanol kills cells...

I’m voting to close this question because it is unclear (e.g. unexplained abbreviations), under researched, and possibly opinion-based.

Check your citations.  How do they use this assay?
I am not familiar with the assay you used, but you are.  Probably it is used to evaluate other plant extracts.  How do those authors do it?   When other people use this assay to look at plant extracts, how do they classify their results?
Consider what has been done in the past using this assay.  Then put your results in the context of what has gone before.


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Why do humans share over 99% of their genomes?
From Wikipedia

The haploid human genome (23 chromosomes) is about 3 billion base pairs long and contains around 30,000 genes.[33] Since every base pair can be coded by 2 bits, this is about 750 megabytes of data. An individual somatic (diploid) cell contains twice this amount, that is, about 6 billion base pairs. Men have fewer than women because the Y chromosome is about 57 million base pairs whereas the X is about 156 million. Since individual genomes vary in sequence by less than 1% from each other, the variations of a given human's genome from a common reference can be losslessly compressed to roughly 4 megabytes.

Why do humans share over 99% of their genomes? Is it a historical accident or a result of selective pressure? If a zygote differed in 5% of the base sequence from the common human genome would it automatically be unviable or could it eventually become a healthy baby?

Dogs vary by as much as 25% and  I think the concencus is interbreeding did happen with other human species, so it could probably be much higher, but not just any genes either.

Humans are a very, very young species, we began diverging on the order of 100,000s of years, compared to e.g. crocodiles which have been around for 200m years, so there has barely been any time for differences to accumulate.

But I'm also not sure this question is well posed. Why wouldn't they share 99% of their genomes?

Let's say there is potential human genome that only shares 70% with the common human genome and that results in you being able to run twice as fast as Usain Bolt. Wouldn't that be evolutionarily advantageous at least in some environments? Wouldn't mutations eventually get us there?

@SKG that's kind of too hypothetical to answer really. If they only shared 70% then they sure wouldn't be human in any sense we know. Evolution happens incrementally, not in massive step changes which would alter 30% of the genome.

I guess if the only option to make embryos is natural reproduction then the question cannot be meaningfully considered. However now that we have several tools for targeted mutations (and also gamma rays for untargeted mutations) I think it does become possible to make an embryo sharing 70% with the common human genome and to put it in a womb. It is probably unethical but I wondered if any expert has written about this.

Relevant: https://biology.stackexchange.com/questions/9172/do-apes-and-humans-share-99-of-dna-or-99-of-genes-what-is-the-difference and also https://biology.stackexchange.com/questions/48084/perplexing-by-the-way-of-estimating-the-shared-or-different-genes-between-humans?rq=1 Make sure when you think and talk about %s of "shared" DNA you know what type of "shared" you mean.

Because the 99% is what makes humans human, of course.  The rest is just minor variation. I would imagine that ANY particular species would have a similar percentage of common genome.  Humans (and other mammals) share a large fraction of DNA with plants, because that's what constructs the basic mechanisms of life, such as cell membranes, ion channels, DNA/RNA replication, &c.

@jamesqf Are there any other species that have 23 chromosome pairs and the same number of base pairs in each chromosome as humans? Otherwise I don't understand how can we compare the genomes of different species (it sounds like comparing strings of different lengths).

@SKG just to give you context, the measured divergence between humans and other great apes is ~1%, and there is complete reproductive divergence between these organisms (https://www.sciencedirect.com/science/article/pii/S0002929707640968). To be clear, these percentages reflect the subset of the genome that is comparable ("alignable") in the first place. Any two humans probably have different numbers of bp in their genomes, due to hypervariable repetitive regions. Moreover, it is relatively easy to blow big holes in the human genome ("aneuploidy"), but it has a tendency to kill the human.

also... "Wouldn't mutations eventually get us there?" i mean... there is a nonzero probability of massive divergence like 30%. After all, evolution does happen on that scale given hundreds of millions of years. But it's like randomly flipping 30% of the bits on your computer and expecting it to keep working. Maybe a very controlled "intelligent design" of bit-flips could get you there, but trial-and-error will take trillions of iterations or more. If you want intelligent design, well, we have to understand the code before we can start hacking! We don't understand genomes that well.

@SKG: As I understand it - and this is not my field, so I'm very far from being an expert - what gets compared are genes.  For instance, there are a family of genes that control the formation of various types of potassiim ion channels.  Each gene is made up of a number of base pairs, and can be located on any chromosome. The particular genes can be isolated and compared: https://en.wikipedia.org/wiki/Potassium_channel

@Armand: You have a couple of undefineds in that experiment.  First, what is the average number of offspring per individual?  Second, what fraction of those mutations result in non-viable offspring?

@Armand tl;dr Your thought experiment is really talking about a different measurement from the OP's cited number. Generally, estimates of substitution rate say ~$1x10^{-8}$ (https://www.nature.com/articles/s41467-017-00323-y), 2 orders of magnitude lower than yours. It doesn't include high mutation rate loci, but those usually aren't alignable anyways so they wouldn't show up in this frequently cited estimate. 1% is probably closer to a "true" measure and ~$1x10^{-6}$ might be closer to a "true" mutation rate, but that is counting a bunch of stuff that people don't tend to count.

@MaximilianPress OK, sorry for the distraction here.

No one seems to be able to decide whether this question is worth answering or not, I will put one down just to bring closure, summarizing some of the discussion in the comments. I think that the question merits answering because the genetics community honestly hasn't done a good job of messaging on this topic.
It's a little involved, so I'll break it into parts. It's a bit long, so for a tl;dr you can skip to the conclusions.
what does "less than 1% different" mean?
First, it's important to know what the widely cited statistic of ~0.1% difference between the average pair of humans means. It means that, of alignable sequence, if you look at ~1000 bp, approximately 1 bp will be different. As a point of comparison, most great apes (including humans) differ from each other by 1-2% by this measure.
However, not all of the human genome is thus "alignable" even between two humans. When I say "alignable", what I mean is that I can take a snippets of two sequences (say, a 100 bp region) and 1) confidently determine that the area is the same on the two genomes, and 2) we can actually associate specific ACGT with each other between the two sequence. Such a 100bp snippet is then "alignable" with regard to the two genomes.
When sequence is not alignable, that usually means that it shows redundancy in the genome, which is to say that it is present in more than one copy, or a similar enough sequence is present that my confidence in a unique placement is low.
In other words, the 0.1% statistic is actually based on the subset of the genome where we can perform confident comparisons. If this minority of the genome were representative, this would not be an issue. But the alignable portion of the genome is highly non-representative, as it contains most of the genes that we care so much about, and none of the big important structural pieces like telomeres and centromeres. This non-alignable region is in part so hard to analyze because it has a much higher mutation rate than the rest of the genome; in other words, we expect the non-alignable regions to be more variable than the alignable regions. Probably much of the genetic variation among humans occurs in these regions.
So the 0.1% estimate is a significant underestimate of divergence. It is nonetheless a useful heuristic for thinking about relative degrees of divergence.
Why don't humans vary genetically more than they do?
This is, I believe, the heart of your question. Whether the true difference is 0.1%, or 1%, or 5%, one could ask why it isn't 30% instead, as you do in your comment. It seems that you are at least somewhat interested in this from an adaptationist point of view, namely that one can imagine that all that extra variation could be helpful in making humans "better", however we happen to define "better" (faster, stronger, smarter, disease-resistant).
This framing is closely related to the classic SE Bio catchall question "Why do some bad traits evolve, and good ones don't?" The common logic being: "I can imagine a way that an organism could be better, why isn't it already better in that way?" I'm not sure that you are specifically interested in that as opposed to a more general curiosity, otherwise it'd be easier to mark your question as a duplicate and move on.
There are several related reasons I can think of why the level of divergence is what it is, I'll try to hit them here:
1. Evolution is slow
Commenter @user438383 got at this a bit in comments, noting that humans are very young in evolutionary terms and asking why not 99%? There's some logic under there that isn't stated that I'll try to unpack.
Other commenter @Armand posted and then deleted (after a too-pointed critique from me) a version of a thought-experiment: let's build a toy model of evolution and plug in some numbers and see what comes out. For example, we could then say that the expected probability of a site remaining the same across $n$ generations is $(1 - \mu)^n$ in a single lineage assuming no back-mutation, where $\mu$ is the mutation rate. We know that the average site in the (alignable) genome has a mutation rate of around $\mu = 1 \times 10^{-8}$. So if we were to calculate that probability for the average site for $n = 100,000$ generations (~3 million years, way older than anatomically ), we get:
$ Pr(no\_change\_at\_position) = (1-10^{-8})^{100000} = 0.9990005$
So, under this deeply simple-minded model with arbitrarily defined numbers, you actually get very little expected change at the position even for rather large time-scales. There are tons of problems with this model, as alluded to by @jamesqf, namely that we ignore population dynamics. But it should at least illustrate the scales that we are working with here.
2. Many/most mutations are shared, due to coalescence
Another fun subtlety is that, of the mutations that do occur in our toy model in (1), many or most of those mutations are expected to be shared between all humans due to coalescence. Briefly, under reasonable assumptions about breeding dynamics, you expect that all humans share a surprisingly recent common ancestor. So there is no variation, because all humans share the mutations.
This issue is exacerbated by so-called population bottlenecks, which are inferred to have happened fairly recently for humans.
3. Most mutations don't matter.
We have known for some time that most of the mutations that do occur have no effect, and allele frequencies suggest no selection occurs on most genetic variation. So there does not seem to be any particular advantage or benefit for humans from having more mutations.
In the absence of selection maintaining a new allele, it is usually lost from the population due to random chance, i.e. "genetic drift".
4. Mutations that matter are mostly bad.
Of the mutations that do have some effect, selection mostly tends to remove them because they are deleterious. These alleles are removed from the population at high frequencies, leading to an incentive to minimize mutation rate to maximize viable offspring.
In fact, the burden of past slightly deleterious mutations retained in the population is a significant force in evolution (the "mutational load"); it is simpler to just not have these mutations in the first place.
Conclusion: you should expect human genomes to differ very little from each other in the alignable regions.
To sum up:

There has not been enough time to introduce all that many mutations into the alignable region of the human population genome.
Of the mutations that occurred, most are shared by all humans.
Of the mutations that occurred, most have no effect, of which many are lost due to genetic drift.
Of the mutations that occurred and had some effect, most were deleterious and were purged by selection.

Did you mean unalignable or "ambiguous location for short-reads"?

mm, I suppose that it's worth clarifying that repeatmasker is the latter, which is definitely an overstatement relative to the more traditional "alignable". I will look around and see if there is a better estimate somewhere.

On second thought, RepeatMasker is likely closer to this than any alternatives. But I can certainly improve the explanation to make it less about short read mapping.

I can't help as I'm just discovering the topic but nanopore website referenced this paper which seems to be exactly doing this, evaluating the true underlying diversity not captured by standard pipelines

@reuns, I'm really thinking of the regions that would not be properly alignable even if we had the perfect full telomere-to-telomere assemblies that the nanopore website refers to. These are significant regions including the telomeres, centromeres, pericentromeres, transposable element sequences, etc. In consideration of your point, I have attempted to reframe the alignability section of my answer to correspond more closely to this, as I was unable to locate a reliable estimate of alignability proportions. It is after all not really important what the exact amount is for this answer.

I think you left out an important point: the differences between humans ARE trivial.  A lot of what goes into making a human also goes into making any primate; much of that is shared with all mammals, mammals share a great deal with other vertbrates, and so on back to LUCA.  For instance, this paper compares the HOX genes (which are fundamental to regulating the body plan) of sharks, various fish, mice, and humans: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC403639/  And then there's what we share with archaea: http://www.sci-news.com/biology/archaeal-chromosomes-07611.html

@jamesqf I think that this is implicit, after all the question could be reframed as "why are the differences between humans so trivial?" That said, I could do a better job making it explicit.


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An apple sunk in water, looks fine. What could it mean?
I washed a dozen of apple just now and one of them sunk in water. Also surprisingly I was not able to find an answer via googling, all top results were about why they float.
The smallest one. I don't know the variety name, but AFAIK it is an apple, tastes like an apple. Could a fact that it sunk mean some internal spoilage?

Image Source: Infovisual: Apple
I think that the density of the endocarp is more than the mesocarp, and in that small apple, more amount of endocarp is present than mesocarp, making its average density more than water, thus, making it sink. Vice versa for the bigger apples.

Welcome to Biology.SE! This looks like it might be a good answer, but answers are much more likely to receive a favorable response if you include supporting references (primary literature is best). Without that support, your answer is indistinguishable from opinion, which is inappropriate for this site. A good example of how to format references.  Please also provide a reference for the image you used. ——— You may also want to take the [tour] and then consult the [help] pages for additional advice on [Answer] effectively on this site.

@tyersome Thank you. I will add supporting references as soon as I find them.


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In vivo, does it take energy to move double bonds from cis to trans in long chain unsaturated fatty acids
In vivo, I understood that because all polyunsaturated fatty acids bonds are -cis they are crowded, kinking the molecule, and because of that, the oxidative enzymes cannot initially access them. They first have to be moved to the -trans side to allow the enzymes to act on the double bonds. So my first question: is ATP needed to move double bonds from -cis to -trans? And also, when the long-chain FA like EPA & DHA are oxidized, because of this energy expenditure to move them, wouldn't they yield less total bond energy when oxidized than if they were just oxidized in vitro, e.g. by heating?
If this is true, then how much ATP is used to first get those double bond groups into the -trans position so the oxidation enzymes can reach them? And … thus could I state that in vivo breaking of long chain polyunsaturated fatty acids would not yield as much energy as would in vitro breaking of these bonds?

As I understand it the question refers to mammalian oxidative degradation of dietary polyunsaturated fatty acids, as the diet is the major source of such fatty acids. Its concern is with two different aspects of this process neither of which involve the use of ATP. The proposition in the final paragraph is therefore incorrect.
The first aspect is not explicitly stated, but seems evident in the mention of the crowding of fatty acids with cis double bonds and the repetition of “in vivo”. I would rephrase it as:

How is the agglomeration of lipids disrupted so as to be available to
  the enzymes that degrade it?

This is as much physiology (not my field) as biochemistry, but there are many accounts available (e.g. this I found at random) regarding the emulsification of lipids by bile salts in the small intestine and their conversion to chylomicrons in the lymph and blood. The latter are either processed and stored in the adipose tissue and later oxidized when needed, or oxidized directly, in the mitochondria, especially of the liver.
The disruption of fatty acid aggregates is purely a physical process facilitated by the chemical environment in which they find themselves. This alone allows oxidative enzymes access. (In passing, I would say that it is difficult to conceive of a mechanism by which ATP could be used to disrupt a hydrophobic aggregate — see Appendix.)
The only stage at which ATP is required is the conversion of free fatty acids to fatty-acyl CoA by fatty acid thiokinase before reaction with carnitine and entry into the mitochondria in the form of acyl carnitine. In relation to the handling of hydrophobic molecules in a hydrophilic environment (or vice versa for membrane transport) it may be worth mentioning in passing that carnitine is a zwitterion.
The second aspect of the question is more straightforward. In chemical terms what the question refers to as “moving double bonds” is called isomerisation. So the question is:

Does cis-trans isomerisation of unsaturated fatty acids require ATP?

This is described in standard biochemistry texts such as Berg et al., from which the diagram below is taken.

No ATP is involved.
Of course a reductase is required to catalyse the reduction of double bonds using NADPH as a reducing agent, regardless of whether they are cis or trans. As the related NADH can be oxidised in the electron transport chain to produce ATP by oxidative phosphorylation, this could be regarded as equivalent to using ATP. However this has nothing to do with the isomerisation.
Appendix: ATP and energy — concepts and  misconceptions
Although this question includes some chemically correct references (the need for cis-trans isomerisation for unsaturated fatty acids to act as a substrate for acyl CoA dehydrogenase), it does raises some suspicions of an incomplete grasp of the factors affecting the viability of biochemical reactions and the role of ATP. If this is correct — and shared by others — I suggest reading an account of how the concept that the Gibbs free energy change of a chemical reaction determines whether or not it proceeds, and how the negative free energy change in the hydrolysis of ATP can be coupled to reactions that have an (otherwise unfavourable) positive free energy change.
Two points relevant to the question are worth mentioning in this regard. Although ATP can be used to cause motion, this involves motor proteins like myosin, kinesin and dynein interacting with large molecular structures. Thermal vibration and thermodynamics are the normal drivers of motion in a cell. The second is that many biochemical isomerisation reactions have standard free energy changes close to zero as, in isolation, the alternative structures have similar energies. What normally prevents the reaction proceeding is the activation energy barrier for the interconversion reaction of one isomer to the other, which is where enzymatic catalysis is comes in. The actual free energy change depends on the concentration of reactants and products, so if the cis isomer builds up and the trans product is removed by reaction then the reaction will proceed in the cis-to-trans direction.


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What’s this fungus growing in supersaturated salt solution
As a hobby, I’ve been trying to “grow” crystals of from supersaturated solutions of aluminium potassium sulphate.
I noticed that my solutions are also rather successful at growing a type of fungus.
The fungus grows bigger and bigger in the saturated solution and leaves behind yellow spores as the salt solution evaporated (see photos).
I’ve always thought that salt is antifungal and anti bacterial, and I’m surprised to see that there is a fungus actually growing bigger in a supersaturated salt solution.
I have tried boiling and filtering (with tea mesh filter so the spores probably got through) the salt solution, but the fungus keeps coming back, so the spores survive boiling too, unless there are spores floating around my room.
Is there a way to identify what sort of fungus it is? What sort of fungus is it likely to be? Have I bred a type of super fungus? Is my life in danger by allowing this to grow?
I live in Sydney, Australia if that helps with identifying the fungus.

It looks like an algae which of course grows very well in sea water. Possibly , in some respects your salt solution is similar to sea salt.

Welcome to Biology.SE! Do you have access to a microscope — I'm pretty sure your question is unanswerable without more information and (assuming sequencing is unavailable) a micrograph would seem to be the best bet. To test @blacksmith37's suggestion, see if keeping your experiments in the dark stops the growth. ——— Please also take the [tour] and then go through the help pages starting with [Ask] questions effectively on this site and [edit] your question accordingly. Thanks!

Thanks, I’ll keep them in the dark to see if that stops the growth.


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Seaweed Farming on the ocean
I discussed with some friends the potential of seaweed farming on the ocean to fight climate change. We asked ourselves how the ecosystem would change when for example seaweed farms are present in the ocean. In a bad and a good way.
What does the ocean ecosystem look like? We know about phytoplankton, zooplankton, whales, fish, sea turtles, and so on.
Do you think seaweed farming on the ocean causes more good than bad to the environment?
Kind regards

Welcome to Biology SE! I'm afraid this question is probably too broad for this site. The ocean ecosystem is too complicated to cover in a SE answer and varies greatly geographically. In terms of sustainability, that will depend entirely on the specific practices used, but regardless I don't see how seaweed farming would sequester carbon. You might be interested in reading about integrated multi-trophic level aquaculture, which is sometimes touted for filtering seawater.

The planktonic algae are of more importance fighting climate change IIRC, there should be some literature around this - I think it's called ocean seeding and basically aims to add minerals to help the algae grow.


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At what point does the capability of sexual reproduction diverge in evolution?
I am asking this question as a layman in biology: What the title is supposed to mean is, as a species evolves, say humans and chimps from its common ancestor, at what point and why are humans or chimps not able to reproduce with it's common ancestor anymore? I'm asking this because two very different looking dogs can still reproduce, so how long would we need to keep breeding dogs until the resulting dogs can't reproduce anymore with the "older" dogs?

Your analogy here is a bit flawed - it is more like current dogs unable to breed with current dogs - which already happens. Very large dogs can't physically breed with very small for instance. The small males can't get up that high physically and the small females can't carry or birth the cross-breed pups as they are too large.

Does this answer your question? How many generations does it take for one species to become a new one?

Zooming out a bit, biologists are usually interested in what practically stops gene flow, not whether gene flow is technically possible or not.  A good example of this is chipmunks on the north and south ridges of the Grand Canyon, USA.  A human could pick up a few chipmunks from the northern side, bring them to the southern side, and they'll breed perfectly fine with the local population.  But in real life, chipmunks cannot naturally cross the canyon and so there is actually no gene flow.
Now to answer your question about when it's literally impossible for two different populations to mix and have children, it's always genetic.  There is simply too much genetic difference between the two species to have a viable offspring. A great example is mules, the offspring of horse and donkeys.  Horses have 64 chromosomes, and donkeys have 63, which is a different number.  For the first generation of mules, it just so happens to not matter. But the dissimilarity prevents mules from breeding and having offspring.  Between any pair of species, there will be genetic differences, and at some point, the genetic differences just become too great to allow reproduction.  For example, you could never cross a horse with a plant because they are simply too incompatible.  However, there is no one compatibility issue that is always responsible.  Different species will have different issues.  It is not always having a different chromosome number.

worth noting chromosome number is not enough several species have different chromosome numbers within the species. like everything in biology its messy and blurry.


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Datasets for Genetic Diversity of Species Across Geographical Distribution
I'm carrying out some research on applying machine learning methods to ecology and conservation. I'd like to know: are there some datasets on genetic diversity across geographical positions? Preferably:

Being well-established and trusted
Including multiple species

Information regarding any similar or related resources will also be highly appreciated. Thanks a bunch in advance, community!

To clarify; Are you asking for datasets that compare differences in genetic diversity between populations within the same species at different geographical sites?

@fileunderwater Not exactly, but I'm so desperate that it would do if there is one :-D

Ok, so what you are after are multi-species summaries of single/point estimates of genetic diversity in the different species, along with geographic location(?). Multiple estimates for each species from different populations are not necessary(?).

@fileunderwater just anything of the kind please

I’m just a bit confused, in the sense that different types of studies and reviews that contain some measure of genetic diversity shouldn’t be too rare or hard to find. If you are looking for more specific types of data, length of study periods or specific methods used that might be another thing. But your question is quite general at the moment. What about this paper or humans and primates; could this be useful to you? Osada, 2015, Genetic diversity in humans and non-human primates and its evolutionary consequences

@fileunderwater Thanks but are there larger datasets?

There probably is, but this isn’t really my area, so I don’t know. Doing some careful literature searches shouldn’t be difficult though. You best bet is probably to do some structured searches to find 5-10 studies of the type I linked and then combining the data from all of them for you training/review study.

@fileunderwater Understood. Thanks!

I'm not really sure what the scope of your research is, but if you are just trying to see if you can successfully apply "machine learning methods" to certain datasets, have you thought about just generating the datasets? Forgive me if I'm making wrong assumptions, but it just seems strange for relevant datasets not to be provided by whoever might interested in gaining real insights on "genetic diversity across geographical positions". Isn't your research overseen or financed by an institution? Just hard to believe that hunting for data would be a more significant part of your study than applying whatever machine learning algorithms are necessary.

Thanks. Sorry I didn't make the scope of my research clear. I want to use methods on REAL data instead of generated ones to evaluate their effectiveness in the task. In other words, the "real insights" you mentioned is the genuinely key part of our research.

Oh and regarding the institution part, our research is funded but they are interested in the methods. So WE need the results for validation, but the funders aren't so much interested in the detailed results of one single test.


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How to quantify basic levels of samples in qPCR?
I do have to quantify the difference in basic levels of expression for a given gene in two types of tissues. There is no treatment, just basic levels.
Before I had a "control" sample that I taken as a "1", then a "treated" sample was calculated based on that control (its values were subtracted), therefore a fold change could be calculated between two types of tissue.
Now there's no fold change, just bare numbers, so I wonder how we suppose to quantify delta-deltaCT ? (what are these references?)

a hint - reference gene... You can easily find the original paper on the d-dCT method with a pubmed search.

if you mean the housekeeping gene - of course, I have it. That is how my delta CT is calculated.

yes, reference = housekeeping. In that case I don't think I understand what your question is. You either do a relative quantitation by d-dCT or an absolute using a standard curve. It looked to me like you were asking to how to do d-dCT.

correct. d-CT is calculated via housekeeping. But when it gets to d-dCT, this is the question. Maybe I just need to compare dCTs between two tissues? This will give me the "raw" number relatively to housekeeping gene.

d-dCT is calculated by comparing your treatment to the control - if you don't have a control then you need to look at your experimental set-up and ask yourself what are you actually looking at (what are you aiming to find out) and what might be an appropriate analysis. This isn't the site to ask those sorts of questions on - check with your supervisor or another student in your lab.


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How to compare basal expression of a gene in two tissues using qPCR techniques?
I have two tissues and they are not treated; I would like just to compare the basal level of expression of the gene of interest (goi) in these two types of tissue (tissue1 and tissue2). After qPCR, I have the following result:

I wonder how am I supposed to make a decision. Will in this case goi be more expressed in tissue2 ?  ctrl - is a housekeeping gene that we use for our types of tissue. Marked in yellow are just mean values. The last column is mean goi_value - mean ctrl_value qPCR quality is not great.

You normalized the data to the control and obtained the
normalized ct-values. Let's call it Δct.
Now you need to find the relative difference between them. In order to do that, the first step is that
you calculate the ΔΔct -> condition_1_normalized - condition_2_normalized.
Calculate the actual difference: condition1/condition2 = 2^-ΔΔct

In your case the answer would be: tissue1/tissue2 = 2^-2.4 = 0.19
The expression of tissue 1 is 0.19 of that in tissue 2. In other words the expression is about 5.2x higher in tissue 2 than in tissue 1.

thank you, but it's not exactly what I need. I am not looking for a fold change, but just a bare data to put it on a dotplot, for example. Say tissue1 must have some value and tissue2 as well. Not that tissue1 is a "control" and considered as "1" or zero. But it must have some value. Both of them.

also how did you calculate that 5.2x ?

I see. If I understand correctly, you want some kind of absolute value for your gene of interest for each of the tissues. Unfortunately, with the data you provided, that is not possible. You can only calculate the relative difference. For absolute quantification you need to run a standard with DNA of known quantity.

The 5.2x is simply the inverse of 0.19 -> 1/0.19
Then instead of tissue1/tissue2 you get tissue2/tissue1.


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What is the best way/kit to extract total proteins from biopsy tissue?
I need to extract total protein from 2mm skin biopsies for western blots.
I was thinking about miRNeasy Micro Kit (50) Cat. #217084 while it's ok for total RNA extraction, I am not sure if it is still fine for proteins. There's a trizol (quiazol) step that provides you with organic fraction that suppose to contain protein fraction, but I am still not sure if it's a best way.
There's also a so-called Urea Sample Buffer (USB), that contains urea, 20% SDS, glycerol, Tris and Pyronin Y - but I am not sure it's a good choice since the percentage of SDS is high and it will be a disaster if I will try to disrupt my tissue.
Or, maybe there's a better solution or kit for that purpose?

This entirely depends on what you want to get out of it. If you need the RNA or DNA, then trizol extraction can work, but no need to buy a special kit for it, just the trizol and chloroform will work too - though for miRNA (as opposed to mRNA), a kit is a good way to go.
If you just want the protein, use a dedicated protein lysis solution for this. Something like RIPA buffer is a good start. make sure you add protease inhibitors and work with cold solutions to prevent proteolysis of the proteins. Depending on what you want to find, you may need to disrupt the biopsies a bit first - chop them into smaller chunks and/or do a collagenase digestion to allow removal of the mass proteins and look at less abundant ones. RIPA buffer is quite strong, so you may want to use a gentler solution without the SDS and/or deoxycholate, but this (again) depends on your final aims.


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How "exactly" is Rabies transmitted?
Context: I know a person which has developed a sort of "phobia" with respect to touching things that has (even the slighest) chance of being in contact with something that can transmit rabies. For instance, if something (a coin) falls on the floor of our university, said person almost cannot take it, since dogs sometimes pass around the place. This has reached a severe level which hinders this person experience in life. However, there is apparently some hope of rationalization. Specifically, the person told me that if someone can prove, for example, that the virus do not "live" long enough after dropping from the saliva of a dog to the floor in order to be transmitted to a human being, then maybe she will be more relieved. This led me to search for information on the internet, but nothing was conclusive. This takes me to my question, and sorry for the sidetracking.

Can someone be infected by rabies by merely taking something from the ground where a infected dog may have passed by? How long does the virus "live" (or is active, I don't know the  proper term) after being "exposed" to the environment? More generally, how exactly does the process of infection happen, and how can it happen?

This is actually far more complex a question than you probably know. The country you are writing from matters. There are some rabies virus variants that have a relatively low pathogenicity for humans (dog rabies is one of these; a relatively large innoculum is required, i.e. a bite, commonly a deep one with lots of virus-laden saliva.) The likelihood of canine-associated rabies developing from a non-bite exposure is extremely low; in fact, the exact risk is unknown.
In the US, enzootic (dog-to-dog) canine rabies virus has been virtually eliminated through vaccination and stray control programs, making wild animals (including feral cats) the primary concern. Depending on the country your friend lives in, dogs may not even be high on the list of worrisome sources.
Regarding canine-associated rabies,

Rabies control was accomplished [through dog vaccination programs, responsible pet ownership, and serious population control of stray dogs] in Europe, North America, Australia, Japan, Taiwan, Malaysia, and Singapore.

If you don't live in one of these countries/continents, dog-associated rabies is still a major concern. If you do live in the mentioned areas, you can probably dip your hand in dog saliva and not be concerned.
Rabies is usually contracted by a bite; that is most common.

The most dangerous and common route of rabies exposure is from the bite of a rabid mammal. An exposure to rabies also might occur when the virus, from saliva or other potentially infectious material (e.g., neural tissue), is introduced into fresh, open cuts in skin or onto mucous membranes (nonbite exposure). ...Exposures to bats deserve special assessment because bats can pose a greater risk for infecting humans under certain circumstances that might be considered inconsequential from a human perspective.
how exactly does the process of infection happen, and how can it happen?

The exact way it happens is not completely understood, but that answer won't help you convince your friend of anything. The virus is usually deposited in tissue near a cutaneous (peripheral) nerve. If it gains entry to the nerve, it makes its way to the brain with the typical disastrous results.
Having said all this, your friend is not going to stop worrying, because the virus is only the focus of an irrational fear. Though the foci of irrational fears are fairly stable, OCD is most likely the underlying problem, and conventional rational discourse doesn't really suffice in curing the obsession.
During 1990-2007, 34 bat-associated human cases of rabies were reported in the US: 6 cases reported a bat bite; 2 cases reported a probable bite; in 15 cases, physical contact was reported (e.g., the removal of a bat from the home or workplace or the presence of a bat in the room where the person had been sleeping), but no bite was documented; and in 11 cases, no bat encounter was reported, but the rabies virus was bat-specific.
Any encounter with a bat, even a dead one (as stated above), must be evaluated for possible post-exposure prophylaxis.
Unfortunately, in the US, animal rabies is common, and at least 23,000 persons/ year receive rabies postexposure prophylaxis (PEP). (It is probably considerably higher, as no reporting mandate exists.) With the elimination of canine rabies virus variants and enzootic transmission among dogs, human rabies is now rare in the United States, with an average of one or two cases occurring annually since 1960.
In the US in 2013, of the three human cases reported, 2 were involved in organ transplants (raccoon rabies virus variant) and one was a Guatemalan (canine rabies virus variant).
In 2012, one human died from an exposure to a bat. He touched a bat under a bridge. He did not report a bite to a witness. He became ill while traveling, and died in Switzerland. A number of humans exposed to his saliva (including his Swiss caregivers) received PEP.
*Some people in high-rabies areas without the illness have been documented to have developed antibodies to rabies. The mechanism is unknown.
Human Rabies Prevention - United States, 2008
Pathogenesis of Rabies
A Single Amino Acid Change in Rabies Virus Glycoprotein Increases Virus Spread and Enhances Virus Pathogenicity
Human Rabies Prevention (Comment From a Canine‐Rabies‐Endemic Region)
Epidemiology of rabies post-exposure prophylaxis—United States of America, 2006–2008
Rabies surveillance in the United States during 2013
Use of a Reduced (4-Dose) Vaccine Schedule for Postexposure Prophylaxis to Prevent Human Rabies: Recommendations of the Advisory Committee on Immunization Practices

First of all, thanks for the answer. I really, really appreciate it : ). We live in Brazil. Does that change or add anything to your answer?

"More generally, how exactly does the process of infection happen, and how can it happen? How long does the virus "live" (or is active, I don't know the proper term) after being "exposed" to the environment?" Good open-access article going over some of the molecular pathology and epidemiology (1). I'd also mention that enveloped viruses are very sensitive to the environment unlike lots of naked viruses, so they don't generally survive long outside the host or get killed easily by detergents and/or desiccation.

@Kendall Is rabies "enveloped"?

@anongoodnurse Also, the person has (as you correctly suppose) been diagnosed by a psychologist as having OCD. Is there anything I can do?

@AloizioMacedo Yes, Rabies virus is a Baltimore V or (-)ssRNA enveloped virus. This basically means there is something similar to a lipid bilayer encasing the nucleocapsid, and the ssRNA has to be transcribed to (+)ssRNA before it can produce it's encoded proteins. The envelope has some advantages in vivo but an exposed enveloped virus is very susceptible to environmental hazards!

@Kendall Thank you, that is very informative. Sorry to be bothering you, but would you mind telling how long is "survive long"? Matter of seconds, minutes?

It varies greatly by virus but, broadly, you can imagine days for enveloped viruses, weeks for naked viruses. There's an open-access article on the matter here.

Let us continue this discussion in chat.

The routes of transmission for rabies are concisely described on the CDC website. To summarize:

The vast majority of cases of rabies are transmitted through the bite of a rabid animal.
For non-bite exposures, there is potential for transmission if an infectious material (such as saliva) enters the eyes, nose, mouth, or wound of a person. There is also potential for aerosol transmission however this is only a risk to laboratory workers.
Petting a rabid animal or contact with blood, urine, or faeces of a rabid animal does not constitute an exposure. (See the CDC guidelines for detailed information on special case contact with certain wildlife species.)

It is possible that if a person touches a fomite (such as a coin) that had very recently contacted the saliva of a rabid animal, they could be infected if that coin came in contact with an open wound or mucous membranes. Neverthless, this is extremely unlikely. Such a case has never been described. Fomites are not considered a route of transmission for rabies.
Furthermore, the rabies virus cannot survive very long in the environment. The virus will die within hours since it is very sensitive to ultraviolet light and desiccation, therefore once the saliva is dry there is no risk of transmission. This process is not instantaneous (the virus does not die as soon as it leaves the host). I was unable to find a precise length of time that the virus can survive in the environment, but this probably has not been researched because there has been no reported cases of transmission via this route. A 1974 study found that the virus in mouse brain tissue died within days at room temperature, and it is plausible that the survival in fluid exposed to the environment would be much shorter.
Most animal workers such as veterinarians receive a pre-exposure rabies vaccine but this is due to the risk of being bitten by a rabid animal.
In summary, we cannot categorically exclude transmission via a fomite, but I cannot stress enough how extremely unlikely exposure to rabies in this manner would be.

This is a good, but not precisely correct answer. Bats are an exception; there have been cases of rabies in people exposed to but not bitten or even scratched by bats. The mechanism is a "postulated unknown bite", but it is postulated. Petting a rabid bat most certainly constitutes an exposure and such a person would be given post-exposure prophyllaxis per CDC guidelines.

@anongoodnurse PEP is not indicated for category I exposures, such as petting a rabid animal - even a bat. It is somewhat dependent upon the history, however. You are correct in that if a person cannot completely exclude the possibility of being bitten by the bat (e.g. they were sleeping in the same room) then they will likely receive PEP. We do err on the side of caution, so I'm sure category I exposures often get PEP, but this has to be balanced against the cost and inconvenience of getting the injections.

Actually, it has to be balanced against the seriousness of the disease; even if the risk is very small, the disease is 99.999999999% fatal. "Cost and inconvenience" may be a concern in underdeveloped nations, but the CDC doesn't consider it in their recommendations. The guidelines are pretty clear; bat encounters are treated very differently (see above).

@anongoodnurse PEP treatment costs several thousand dollars, so of course cost is a consideration. Of course the severity of the disease is the major factor, which is why I said we err on the side of caution with treating. For category I contact where there is an extremely unlikely risk of transmission (no bite, no scratches, etc) then the CDC would not always recommend treatment as you suggest. In this answer, I am just trying to convey that the risk is never 0, but extremely low.

I don't like to debate in comments. I don't know how decisions are made in your country, but I'm familiar with how they are made in the US. I am fully aware of CDC recommendations (as well as regional and state department of health recommendations) and I disagree with you. Furthermore, you might find the fifth reference interesting; it discusses the very issue.

Let us continue this discussion in chat.


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What are some useful (starter) metrics to use on phylogenetic trees?
I'm doing a computational biology project in which I simulate evolution under different inheritance rulesets and I am generating phylogenetic trees (beautifully visualised in Python with ete3).
My question is: where can I find and test out some simple metrics that can describe these trees in terms of 'branchy-ness' (you can tell I'm not a bioinformatician or phylogeneticist!). I'm looking for kind of mean-field descriptors of the trees. Kind of like degree distribution for networks.
My specific goal is, like in network science, trying to see if I can differentiate between different rulesets that created the tree by examining the [statistical] properties of the tree itself.

Is there any specific goal you want to achieve by comparing these metrics between different trees? You could count the number of branches, time to the common ancestor of all represented lineages, the total quantity of "evolutionary time" as the total length of all branches, the mean speciation time, ...

The specific goal is, like in network science, trying to see if I can differentiate between different rulesets that created the tree by examining the [statistical] properties of the tree itself.

Here are a few metrics you can calculate to get you started. You can use R to perform these calculations.
Net Diversification Rate (r)
Net diversification rate is (rate of speciation - rate of extinction). You can calculate it using the bd.ms or bd.km functions in the geiger package for R.
r = 1:

r = 2:

Tree Imbalance: Colless index (I)
Colless's index measures how imbalanced the tree is. It sums the differences in the number of clades in each pair of taxa. You can calculate it using the colless function in the apTreeshape package for R.
I = 0

I = 21

Branching Times (gamma)
The gamma statistic measures how much later or earlier branching times happen than what you would expect from a normal birth-death process. A negative value means branching times occur earlier, positive means branching times occur later. You can measure gamma (and generate a lineage through time (ltt) plot, as shown below) using the ltt function in the R package phytools
(From this Nature article)

These are just a few of many parameters you could calculate for a phylogenetic tree. It should be enough to get you started.

awesome!  thanks a bunch. i'm using python, so hoping I can find similar packages to measure these things there (maybe in ete3 ? )

Call R from Python: http://rpy.sourceforge.net/rpy2/doc-dev/html/introduction.html

cheers for that. i'll check it out.


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Can whales send each other [3D] images?
Russell Arnott, apparently an oceanographer, who currently works as a project manager at the Sainsbury Laboratory, Cambridge University (UK), wrote in his piece Whales and Waves (2016, p.2):

New research from the DAREWIN Institute postulates that once a whale has obtained an image of an object using sound, it can then send a holographic image of what they saw to another whale using sound.  That’s like sending a 3D-picture to your friend using only your face.

Although that article is available on Academia.edu, and Russell Arnott does indeed apparently currently work for Cambridge University, the article is not from a published academic journal. With my meagre understanding of biology I've tried to hunt down articles on this in peer reviewed journals or academic books but have drawn a blank.
Is there any research which suggests that whales may be able to send each other three-dimensional images, or in fact, images of any sort at all?

whales uses sonar and it might be possible for other whales to see the return signal/picture,it is sort of scary if it is done in any other way than this.

Computers transmit topographic maps, 2D images and 3D files with colors and measurements at high bandwidths, Animals transmit symbols as language. If you mix the notions of 3D and symbols, it's like saying the whale's don't need to define the seabed or a shoal of fish when they speak, they provide a high bandwidth topographic map, and that's a muddled suggestion without data. Given their soundwaves and the precision of analogue time measurement, they could speak all day to transmit only a small topographic scan.

To send a holographic image using speech, you have to convert the information of the image from an echogram to vocal soundwave and back to an echogram, using analogue time measurement process the data. for a 3D image, whales also have to take multiple echograms from different locations, think about them, then synthesize them into one low bandwidth voica call going through a different sensory pathway, using analogue time for precision, and no mammals have very precise timing to "send images" using sound.

@bandybabboon Apart from humans animals don't use language. And my interest in this question is precisely because it throws light on why humans need language to communicate and why we have verbs.

Holographic is a very uncommon word to use for animal communications. 3D scenes definitely require adding together more than one echogram.

Dolphins, primates, prairy dogs, elephants and many other animals communicate symbols, maybe called signalling systems not language.

As somebody familiar with production of the first digital 3D images, Russell Amott's suggestion of whales communicating in image format seems entirely possible.  Perhaps the people at CETI (organization studying whale communication) need a few top geophysicists to consider the problem of how to decode whale language?  Why not try running whale communications through seismic data processing algorithms?
Consider that a whale spends a large, continuous amount of time decoding sonar clicks into a 3D spatial understanding of its surroundings.  Sonar works by measuring the phase shift in sound REFLECTIONS.  Consider that in order to produce the loudest sounds it modulates sound also through internal reflections within its own biology (i.e. "junk" of a sperm whale).  Is it not reasonable to consider that a whale would evolve to produce sounds analogous to what it hears and decodes?
This would be a great problem to get some interested Geophysicists to attempt to solve.  I believe this is a more probable manner in which to decode whale communication than trying to use purely human language models.
Imagine if you could transfer 3D images into somebody else's head?   No need to go to the movies.  Wow.  Maybe that's why a sperm whale's brain is six times larger than a human's?

I'm confused why this is getting any upvotes. There is no citation of any relevant information here, purely speculation, and it is simply not physically possible to transmit echolocation sounds the way they are heard, because they depend on arrival times at the listeners ears; regardless of what you hear, you can't transmit without separate speakers that can control the arrival time at each ear and also encode spectral cues that are distinct by the individual listener, only meaningful to that listener, and applied to any sound that comes in according to the direction of that sound.

@BryanKrause But I can hear someone singing and receive that signal through two ears at different times mediated through my own personal biological equipment and still transmit version of what I've heard with meaningful accuracy. The sound that's coming back to a whale/porpoise/dolphin originated from that mammal from roughly the same area as it's received (and the reception isn't anyhow simply directly through the ear , but is also transmitted through fat in the jaw to the ear). The buzzes going outwards are made by the mammal itself. It is not unreasonable to suspect ...

@BryanKrause ... that the incoming signal can also be reproduced. If what is being received by one mammal is reproduced to another, it does not mean that that other mammal will not be able to understand that the original receiver is transmitting what they 'heard' fro their own point of view. I also do not see that two ears are necessary for decoding the original signal to reproduce a three dimensional representation. One might do adequately on its own.

Just because a person shines a torch and others see what the light hits and reflects off doesn't imply communication. @Araucaria Whilst I see that this answer touches upon unknown neural and conceptual processes that occur (or may occur) in whales, it doesn't seem to answer the question about communication, nor is it supported with even minimal references.

@JiminyCricket. That isn't a good description of what I described, which involved an animal resending the signal they received - handing over a polaroid in your analogy. The 'communication' seems to have been imported into the discussion by you! However, as a relevance theorist working on human linguistic communication, all that would be needed would be for the sender to be ostentatiously and self-consciously trying to affect the receiver's stock of assumptions about the world, and for the receiver to be aware f this intention, and for the receivers recognition to be understood by the sender.

@Araucaria Read more carefully what I've said and think about how the spatial representation of the world relates to the spatial representation of the ears. You cannot deliver that signal because it needs to come from all the real physical places at once. You can't generate a mimic of it from one place, it's not physically possible,  because sound travels freely: the right ear is going to receive the signal you intended for the left and vice versa.

I have direct knowledge of programming 3D images and know the maths processes inside out. digital is far less useful than analogue, that's why humans don't transmit numbers to each other when they describe scenes, and whales deffo wouldnt either unless they favor low resolution and efficiency.

This just seems like a weird and potentially hyperbolic way to state a postulate that whales use language to convey things to one another that they have observed via echolocation. Don't overthink it.

There are currently no serious claims of that type by any scientists who use the scientific method.
If it's an inaccessible, unevidenced, unpublished, unreviewed, unauthored claim of a theory, that nobody has written about from the science community, publicly attributed to no researcher titles, you can presume it is not real.
1/ The linked article quote "is for an audience of 15 years old average GCSE Students."
2/ "DAREWIN" 3D, i could find zero results related to any related research. No data is available on that theory from DAREWIN. Please comment if you find anything else than that one single statement meant for a school project.
I know about the TF dimensional limitations of sound. The article suggests that the whale sounds work like sonic modem rather than a language.
I programmed 3D graphics, vectors, maths and geometry for more than 5 years and sound waves for 8 years, including dolphin TimeFreq graphs, data transmission using sound clicks.


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Do mycorrhiza only form with angiosperm?
if not than with which Phylum it forms

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Mycorrhiza refers to any fungi that forms a symbiotic relationship with a plant. There are many different species, and they are necessary for healthy plant growth in ~70%-80% of vascular plant species with representatives from more than ~90% of plant families. Needless to say, mycorrhiza associations with plants are very common.
What you may be thinking of is rhizobia, which are not as ubiquitous.  Rhyzobia are soil bacteria associated with legumes (peas, soybeans, clover, lentils, etc.). Rhizobia fix nitrogen for the legume, and the legume forms root nodules where the rhizobia live. The nodules provide an ideal location for the rhizobia to live and thrive. This relationship has only been observed in legumes (scientific name: Fabaceae).
In summary:
Both mycorrhiza and rhizobia form Symbiotic relationships with plants. Mycorrhiza are fungus that form relationships with most species of plants. Rhizobia are bacteria that form relationships only with plants in the legume family.


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Methods to obtain the mass distribution of an ensemble of bacteria?
Are there any mainstream methods to obtain the mass distribution of a bacterial population?
What kind of assumptions to they make?
Could anyone point me to some references?

Well, careful centrifugation can separate cells based on size.

Short Answer:
Flow Cytometry or Dynamic Light Scattering (DLS) might be the most "mainstream" methods out there that might give you a size or volume distribution. Then you need to do some reading how to get bacterial mass from size or volume.
Image Cytometry:
Krößbacher et al. 1998 used electron microscopy (which is not mainstream)
Oldewurtel et al 2021 used Phase microscopy and complex image pipelines (also not mainstream, but definitely cheaper than electron microscopy). They also stress that "surface-to-mass ratio remains nearly constant", which is very nice, since that means that one does not need a weighing scale, but only know the size of bacterial particles.

Therefore, cellular dry-mass density varies systematically with cell shape, both during the cell cycle or after nutrient shifts, while the surface-to-mass ratio remains nearly constant on the generation time scale.

Flow Cytometry:
Knowing that one can determine bacterial mass from its size, one might just use flow cytometry to get the size distribution (see Felip et al. 2007) and then calculate the mass.
Dynamic Light Scattering
DLS can also determine size distributions of particles and was used with bacteria (Vargas et al. 2017).
(Field-Flow Fractionation)
Something fancy I found: Sharma et al 1993

Thanks, and how do people estimate the density of bacteria? (needed to go from a size distribution to a mass one)

@Gyromagnetic: Do you really need the exact mass distribution (as opposed to a relative mass distribution)? Either way, you can here use macroscopic measurements, looking at the average volume and mass of a bunch of cells.

In addition to Markur’s answer, Coulter counters suck the cells through a small capillary (in an electrolyte solution), which also serves as a conductor in an electrical circuit. They then estimate the cell volume by the change of conductivity of the capillary.
In my experience this method is more easy and robust than many others (e.g., flow cytometry) and as far as I can tell, the device is rather cheap and low maintenance – on account of being established and mass-produced technology.
Some relevant assumptions that go into it are:

If you care about relative volumes: All cells are equally affected by the electrolyte solution and have the same conductance properties. If you care about absolute volumes: The cells are not affected or in a known manner.

Cells do not stick to each other in a way that prevents disentangling double measurements, i.e., two cells entering the capillary at the same time. (If this happens by pure coincidence, you can statistically handle this.)


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How to obtain raphides from plants?
I recently found out that certain plants produce needle-shaped crystals called raphides. I am a physicist and I have an experiment in mind that could use these needles, but I would need to extract them from the plant and keep them in a liquid solution somehow.
How can I do this? Is this possible/hard to do? Does anybody know how to do this or have any experience?

Searching for raphide purification I came up with a paper where they isolate these crystals from kiwi fruit:
Konno, K., Inoue, T. A., & Nakamura, M. (2014). Synergistic defensive function of raphides and protease through the needle effect. PloS one, 9(3).
Does that help?
Note, however, that while according to this paper these large crystals can be kept in suspension that isn't the same as a solution ...