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Why can't diamine oxidase be supplemented?
I've read from an academic article that diamine oxidase cannot be supplemented  but it had no explanation as for why. I am curious as to why this?

Food rich in histamine or red wine may cause allergy‐like symptoms
  such as sneezing, flush, skin itching, diarrhoea and even shortness of
  breath. The suspected reason is a diminished histamine degradation
  based on a deficiency of diamine oxidase. As diamine oxidase cannot be
  supplemented, a histamine‐free diet was implemented to reduce
  histamine intake.

Proteins other than hormones in general are not supplementable. Think of genetic retinitis pigmentosa, metabolic diseases etc. They are digested when taken orally and won't reach intracellular targets when injected in the bloodstream or in tissues.

More recent research contradicts the statement that diamine oxidase (DAO) cannot be supplemented1.
This paper clearly supports use of DAO. However, Triptans like sumatriptans are typically used instead because of cost.
Note: Research is of 2018, so not your fault.
Reference:

Izquierdo-Casas, J., Comas-Basté, O., Latorre-Moratalla, M. L., Lorente-Gascón, M., Duelo, A., Soler-Singla, L., & Vidal-Carou, M. C. (2019). Diamine oxidase (DAO) supplement reduces headache in episodic migraine patients with DAO deficiency: A randomized double-blind trial. Clinical Nutrition, 38(1), 152-158.

The OP's linked article clearly states that DAO can't be supplemented. I'm guessing that what you mean by "The article" is the one you linked? I've edited based on that assumption, but please correct as needed. Also, can you elaborate on the statement about triptans — it isn't clear to me how that is relevant to the original question.


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Is NADPH unstable in UV light?
I am working on an enzyme activity assay using NADPH as a cofactor. The MSDS for NADPH Tetrasodium Salt, tells to store it in a place away from heat and light. Does this include UV light or just visible light? I was told that a pure NADPH stock solution (with pH 7 phosphate buffer as solvent) could be re-used after absorbing UV light at 340 nm, but based on my understanding of the MSDS it seems that shouldn't be the case. So my question is: could I reuse a sample of pure NADPH stock solution that was exposed to UV for my enzyme activity assay?

In my experience, reuse of NADPH stock solutions maintained at pH 7.0 or above is perfectly acceptable. Taking an absorbance spectrum every half-hour or so will probably show that definitively.  However, NADPH is VERY unstable in acid, leading to a 'bleaching' of the absorbance at 340nm.  Try measuring a rate at pH  6 or below, 340nm,  with 100 micromolar NADPH and no enzyme or cosubstrate. There will be a very significant decrease in absorbance with time (ie a 'blank' rate) which will make estimation of enzymic reactions at low pH virtually impossible.

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Evolution of endosymbionts?
Mitochondria and plastids in eukaryotes evolved through a process of endosymbiosis. How does an event like a eukaryote engulfing a bacteria, become a part of the genome? Some of these primitive eukaryotes could have been more predisposed to more vigorous endocytosis as a result of their DNA, but how could the physical event account for evolution of organelles?

What exactly are you asking here? Mitochondria still have their own DNA, it's not in the "parent" cell. They reproduce on their own, it's just that they wait for the cell reproduction signals to do so. In fact, this is exploited in mapping female genealogies, since mitochondrial DNA is transferred predominantly through the female line (egg cells tend to carry a lot more mitochondria than sperm cells).

If your question is about how the engulfed endosymbionts lost their genes and became dependent on the host cell then this is the answer.
The current theory (Limited Transfer Window hypothesis) says that at some point, injury or membrane rupture of the endosymbionts would have resulted in the leakage of their genomic DNA. The host genome would have acquired these genes by a lateral gene transfer mechanism. Now, because the endosymbionts that have lost these genes randomly, would have a lower cost of maintenance (as they can acquire these gene products from the host cell) compared to those who still maintain these genes themselves, the former will outgrow the latter at some point. Note that there is a constraint on the population size and therefore there is a good chance that the endosymbionts with full set of genes, could go extinct because of their additional growth cost.
This is a probable way by which the endosymbionts lost their genes and became dependent on the host. It has been observed the number of genes transferred to the host correlates with the number of endosymbionts present in the cell. The rationale behind the limited transfer window hypothesis is that if the number of endosymbionts is high, then the likelihood for the host to tolerate damage to a few endosymbionts is higher (note that damage was the first step in the endosymbiont to host gene transfer).
This is still a hypothesis and the exact mechanisms are yet to be elucidated.

References:

Timmis, Jeremy N., et al. "Endosymbiotic gene transfer: organelle
genomes forge eukaryotic chromosomes." Nature Reviews Genetics
5.2 (2004): 123-135.
Smith, David Roy, Kate Crosby, and Robert W. Lee. "Correlation between nuclear plastid DNA abundance and plastid number supports the limited transfer window hypothesis." Genome biology and evolution 3 (2011): 365-371.
Lane, Nick. "Plastids, genomes, and the probability of gene transfer." Genome biology and evolution 3 (2011): 372-374.
Gray, Michael W. "Mitochondrial evolution." Cold Spring Harbor perspectives in biology 4.9 (2012): a011403.

Thanks for the information. I have a great deal more information than when I started. Rupture of the membrane could explain acquisition of the foreign DNA.

@user17496 But you should bear in mind that it is just a hypothesis. This has not be explicitly proven.

@WYSIWYG Once the DNA is out of the symbiont, it would likely be like transfecting a Eukaryotic cell today, easier as the symbiont is already in the cytoplasm. The chromosome would need to get into the nucleus. It would need to be linearized, and there would need to be an area of complementarity that would allow for strand invasion. The rest would be handled by the cells own repair mechanisms. Genes are maintained through selective pressure, and as you said, if it costs less to have the product produced by the host, then sustaining deleterious mutations will not be under selective pressure.

@AMR Complementarity is not really essential. They can insert via a break and join mechanism. But as I said, we really do not know the mechanism yet. I don't think even the insertion hostpots have been studied in great detail. That reminds me; I should look for that.

No, not essential, but I was thinking along the lines that there would be more opportunity for homologous incorporation than going down the path of NHEJ, especially to get a productive insertion where the gene is downstream of an active promoter, unless the entire gene "cassette" was inserted. Anyhow, my crystal ball gets very fuzzy after looking about a billion years back.

I think you mean mitochondria and chloroplasts and are referring to the endosymbiont theory (Reference to the unedited question as it was originally asked).
The definition of symbiosis for the most part tells you how coevolution can be accounted for by this theory.
The larger Eukaryotes provided the protomitochondria with a protected environment for growth and the protomitochondria produced usable nutrients for their hosts from molecules that the eukaryotes were unable to process on their own.
The two organisms struck up a balance and the mutually beneficial relationship allowed both to prosper. As the Eukaryotes with protomitochondria were better able to utilize more of the chemicals from their environment than those that did not play host to the protomitochondria, they had a survival advantage that other early Eukaryotes did not.
As plant cells have both mitochondria and chloroplasts, it is believed that one of the eukaryotes with a mitochondria endocytosed a photosynthetic prokaryote. This event wasn't as much of an advantage as the first symbiotic relationship. Since all eukaryotes have mitochondria or evidence of ancestors that had mitochondria, but not all have chloroplasts, then it was beneficial enough that the photosynthetic organisms were also able to coevolve within the eukaryotes and alongside the mitochondria, but they were not beneficial enough so as to give those cells that hosted chloroplasts enough of an advantage that they took over all life on Earth.
Mitochondria also have their own DNA, so their complete genome is not found in the eukaryotic genome. Over evolutionary time, mitochondria have lost the genes that allowed them to live freely, so you could not extract them from eukaryotic cells today and expect them to grow external to the eukaryotic cell. They require substances that the eukaryotic cell makes. Whether there was recombination between the two genomes or just that both had the same function at one point and then through genetic drift caused the mitochondria to lose the homologous gene, I could not say.

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What is the antagonist of epinephrine?
Norepinephrine and epinephrine have similar stimulant properties on the nervous system and the body. I was wondering which hormone is the antagonist of epinephrine?

You should understand that it is not the neurotransmitter that is responsible for an effect; it is the neural network that causes an effect and the neurotransmitter is just a mediator.
In CNS, glutamate is generally used in excitatory synapses whereas GABA is chiefly used in inhibitory synapses. Similarly, in peripheral nervous system epinephrine and acetylcholine are used in excitatory synapses whereas glycine is used in inhibitory synapses. Some neurotransmitters always cause excitation in the post-synaptic cell (for example glutamate), however, they can also cause activation of inhibitory neurons and thereby cause inactivation, indirectly. Also, the post-synaptic effect depends on the type of receptor expressed by the post synaptic cell and not the neurotransmitter per-se. For example, α-adrenergic receptors cause smooth muscle contraction whereas the β2- receptors cause smooth muscle relaxation [1]. Similarly, there are both inhibitory and excitatory dopamine receptors [2].
The two divisions of the autonomic nervous system — the sympathetic and parasympathetic systems act in a complementary manner. The former uses epinephrine and norepinephrine whereas the latter uses acetylcholine.

You hit the nail on the head +1


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What are possible reasons for a DNA template not giving amplification?
I am trying to amplify ~10kb length fragments using NEB LongAmp Master Mix. I had been using one particular aliquot of DNA (template amount in each PCR reaction of 10ul being around 60ng) and getting amplification results as desired. However, when I switched to a different aliquot of similar concentration, I am not getting amplification all of a sudden. I tried a different DNA sample and different primers and got results. So the problem lies with either the template or the primers. I have tried increasing amount of template, increasing Mg2+ by 1mM in the final reaction etc. but to no avail. The DNA templates/100x primer stocks are stored at -20⁰C and the DNA had been phenol-chloroform extracted. What could possibly be wrong here ?

When you changed only the DNA you use it is pretty likely the reason. How do you prepare this DNA? Was there anything different than in the batch before?

No. I had extracted it using Phenol Chloroform method and had the original DNA concentration at 700ng/ul. From this, I used 1 part DNA and 9 parts RNA free water dilutions to use for PCR. When I made a new batch of such dilutions recently, I am not getting any amplification.

A new batch of dilution or a new batch of DNA which was diluted this way? From what tissue or cells do you extract the DNA?

Possible problems:

Degraded primers: What were they dissolved in. Run a 10% PAGE in 0.5x TBE to see if they are fine
Degraded template: Check on gel. 100ng is sufficient to see on the gel.

When doing the phenol chloroform extraction, are you doing a chloroform only extraction to remove leftover phenol? Are you sure you're only taking the aqueous layer? If you're pulling phenol or chloroform into your aliquot of DNA those could interfere with the enzymes.

Lets summarize the comments into a real answer: When the only chemical changed in your reaction is the DNA, which came from a fresh Phenol/Chloroform preparation, I would suspect it. From there, we have a few possibilities, what could have gone wrong.

The most likely cause of an enzymatic reaction not working with
Phenol/Chloroform prepared DNA are left-overs (or carry-over) of the
organic solvents from the reaction. The inhibit (especially Phenol)
enzymatic reactions very efficiently. It would help to do another
round of DNA precipitation.
It is possible that your DNA didn't got dissolved after the drying
step in the precipitation. Depending on the conditions, DNA can form
a pellet which is hardly to see and even harder to dissolve. So your
DNA concentration would be too low to get amplification. To prevent
this measure the concentration of your DNA.
It is possible that the water you used is contanminated with DNAses
(although it shouldn't) and your DNA got degraded by them. Here only
new aliquots will help.
Depending on the size of your amplicon, it is possible that overly
harsh condition during the preparation led to the shearing of the
DNA leaving not enough intact DNA to get a proper amplification. If
this is the case, be more gentle with the extraction.

Additionally your primers can be a problem. It would be good to keep the 100x stock solutions permanently at -20°C and prepare 10x working solutions. These can undergo freezing and thawing (which will degrade them over time) but they can be replaced easily.
I would also try the new DNA prep (if possible side by side with one that is working) with new primers to test if they are the problem.


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SPR TRIS or PBS buffer
Hey guys I want to ask why I can't get any binding interaction if I use TRIS-NaCl buffer for protein quantification using aptamer as its bioreceptor. But I can see the binding interaction if I use PBS buffer
The procedure step for my experiment :
1.) Self Assembled Monolayer (24-48 hrs) - NHS/EDC Activation (wash with acetate) - Streptavidin (Acetate buffer) - wash with acetate - wash with TRIS - ETH Blocking - wash with TRIS - Aptamer biotinylated (TRIS buffer) - Protein (TRIS buffer)
all washing buffer are using TRIS or PBS
And down below (my other method) I tried to exchange the buffer from PBS to TRIS after aptamer injection
2.) Self Assembled Monolayer (24-48 hrs) - NHS/EDC Activation (wash with acetate) - Streptavidin (Acetate buffer) - wash with acetate - wash with PBS - ETH Blocking - wash with PBS - Aptamer biotinylated (PBS buffer) - Buffer exchange (PBS to TRIS) - Protein (TRIS buffer)
I can see my aptamer signal in 1st and 2nd method, but still no binding occurred between my aptamer and my protein in TRIS buffer.

I don’t quite understand your protocol, but EDC/NHS activated carboxylates react with primary amines. Tris contains a primary amine and will be present at a much higher concentration compared to streptavidin.

Are the pH and ionic strength of the two buffers the same?

I presume the aptamer is biotinylated; if not, that’s a likely reason for the outcome.
Have you used a biotinylated positive control to confirm the surface is functional with PBS? I understand you want to buffer exchange to Tris, but this positive control in PBS buffer is important.
Amine coupling involves covalent attachment of the 1-Ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC) to carboxylate on the chip surface, followed by replacement of this adduct with N-hydroxysuccinimide (NHS), thus creating an active surface to crosslink primary amines at slightly acidic pH. Then the unreacted active surface is quenched with ETH. Residual active surface would react with Tris that contains a primary amine.
Perhaps the ethanolamine blocking agent may not have completely inactivated the active surface following streptavidin amine coupling, enabling crosslinking of Tris buffer to that surface and impacting the ‘Protein’ capture?
Other possible reasons: Binding of the aptamer to streptavidin may sterically hinder binding of the 'Protein.' Also, the loss of the proper secondary structure of the aptamer may impact the 'Protein' binding. Perhaps refolding the aptamer prior to streptavidin capture may help?
Edit: Differences in pH and ionic strength may impact 'Protein' binding to aptamer when tested in Tris versus PBS buffers.

Yes, the aptamer is biotinylated thus it can bind with the streptavidin. Yes and I also know that NHS ester will be released because of the primary amine of the TRIS buffer and create covalent amide bond.
But I'm pretty sure right now that my ethanolamine blocking agent is deactivated all the NHS ester residue because I also double-check using BSA blocking to my SPR surface. That's why I'm pretty confused why with PBS my aptamer can catch the protein and with TRIS can't

It looks like the ethanolamine quenching may not have been sufficiently effective?

Oh I forgot to mention that I can see my aptamer signal, but when I inject my protein I can't see its binding interaction and the signal keep going flat (not indicating any changes at all)

OK. Increased RU counts is a good sign. Has this lot of protein been confirmed to bind this lot of aptamer by a different technique?

I got this aptamer and its protein from my co-worker and he said he already test and confirm it with ELISA.

But why I can see the binding interaction of my protein and its aptamer using PBS buffer?

I presume you have a control surface with no streptavidin, but everything else to confirm the ethanolamine quenched the residual active surface? Also, is the positive binding of the protein to its aptamer using PBS show reasonable dissociation rate constants?

Yes, and i see reasonable association and dissociation rate constants of different kind of concentrations

At what pH was the 'Protein' binding done in the PBS and Tris experiments?

Ah yes maybe because the pH and ionic strength difference, so because of that my aptamer cant bind with my protein properly even though my aptamer can bind with streptavidin.

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

Can you please expand your acronyms?

@jakebeal sure!

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Can typed-switched B cells recognize the tertiary structure of antigens?
As T helper cells are responsible for isotype switching but Th cells are MHC class 2 restricted and MHC only presents denatured peptides (up to 30 residues, no folding), does that imply that only IgM can recognize tertiary structure? If IgG and the like can, how was that naive b-cell able to present a tertiary structure recognizable by a Th and thus typed-switched.

MHC and B cell receptors/antibodies are totally different things, and recognize completely independent antigens. I think you're mixing up the two.

But doesn't it take a Th cell specific to the same antibody as a B cell to cause the B cell to type switch? And the Th is immune restricted to class II MHC.

Th cells are not "specific to the same antibody" as B cells because T cells don't care about antibody/BCR recognition sites. B cells are activated by Th cells that recognize the same antigen (a protein, for example), but not the same epitope. BCRs/antibodies and MHC (I or II) have very different epitopes. A B cells Class II doesn't necessarily display the same peptide its BCR recognizes.

Sorry, meant antigen, typo.. how does the Th cell know it recognizes the same antigen as the B cell (or vice versa)?

Ah, didn't realize that b cells endocytos BCR-bound antigens and present on MHC class 2.. makes sense.

B cells endocytos BCR-bound antigens and present them on class II MHC. Previously activated Th cells then activate the B cell.

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Does common PCR amplify genes regardless of what cells / barriers they are in?
I have some understanding of how PCR testing works. What I have always been wondering: how can we be sure that a primer reacts with the targeted gene(s) regardless of where¹ the genes are inside a sample?
¹: Specifically with "where" I mean what cells or other barriers they are inside. In other terms my question is: how can we be sure primers reach every possible target?
In more detail: when targeting a gene that is part of the host organism's own genome things might be quite clear, we can have strong assumptions of "where" the gene is. But when targeting genes that are part of organisms that have a parasitic relationship with the host, a very complex system of  interactions (mostly called "immune system") should govern the (co)existence of host and parasite. These complex interactions might lead to situations in which the parasitic organism is present in a very special, isolated manner / place in the host. To give one example: micro organisms in the cell walls of macrophages that are about to undergo lysis. Can we be sure that common PCR would detect genes of such micro organisms? Why?
EDIT: as pointed out in the comments the perhaps most correct phrasing is, how do the DNA isolation techniques employed guarantee that primers reach every possible target?

PCR isn't conducted inside living organisms. The DNA must be isolated first.

That is correct and was clear to me when posting the question. So my question based on your comment also could be phrased: how do DNA isolation techniques guarantee that all possible targets' DNA is isolated from a sample. I shall update my question with your more precise phrasing, thanks!

You will only detect the DNA that you put specific primers in for. You could use random primers, but then you will have a hard time quantifying what you are amplifying. If you are doing Transcriptome analysis in Eukaryotes then you can us a Poly-T primer and Reverse Transcriptase to take mRNA to cDNA. From there you would need to run some form of microarray to detect what it is that you are pulling down. So, no, unless you are putting primers in for specificity for the microorganism you have, you should not get amplification, unless you use random primers.

@canadianer That is false. Colony PCR can allow amplification of bacterial DNA even when the bacteria are added into the PCR mixture while still alive and viable, and the same applies to mammalian cell culture.

@MarchHo Nowhere in that description do I see PCR conducted inside living organisms.

@MarchHo canadianer is basically correct: the KIT you have linked uses denaturation of proteins at mostly 95°C as described in the instructions. Thus any organism whose DNA is isolated is dissolved. You are right the microorganisms and mammalian cells are alive when added, but they are decomposed during the process exactly the way canadianer explained.

While your understanding is correct, I still disagree with @canadianer 's phrasing. The DNA is released from the cells, but at no point in time was the DNA ever purified or isolated prior to the PCR reaction's completion.

There are many methods of DNA isolation, so it is difficult to make broad statements about all situations. Typically, though, during the DNA isolation protocol, essentially all proteins are denatured and removed, as is RNA, cell membrane components, extracellular matrix, etc. If you are using a high-quality well-validated kit from a reputable supplier, your final product should be just DNA. In your hypothetical scenario where parasites have been engulfed by macrophages (BTW, eukaryotes don't have cell walls), as long as the parasitic DNA hasn't been degraded, it will be present at some level after the purification process. As long as it is above the minimum sensitivity of the PCR reaction, any parasitic genes that are being targeted will be amplified.
During the PCR process, the heating step to 95°C completely denatures and melts all the DNA present, allowing the primers unrestricted access to their complementary sequences, where they can then base-pair during the annealing and extension step.

Thanks for answering! Can you explain in more detail how the DNA from a microorganism engulfed in the cell membrane of a macrophage will make it through the isolation process? Why does high-speed centrifugation not discard the macrophage entirely? Can denaturation (or another process) as employed in isolation techniques decompose a macrophage's lipid membrane and the engulfed microorganism's cell wall?

@BM there are different kits for different types of samples, so if you're processing samples that have cell walls, make sure you choose the correct reagents. The lysis process is quite harsh, denaturing pretty much everything. Look through some of the information at Qiagen, one of the most highly-regarded companies in this field. The centrifugation steps are after lysis (the other answer is rather confusing in this regard), so you're not losing any cells.

@BM Here is a selection guide for what type of kit to choose for purifying genomic DNA.

Thanks for the link and your comment. I see that the whole issue is more complex than I thought and boils down to what isolation KIT you use. I still want to know much more precisely how those work however, so I will accept your answer and open a new question with more details.

@BM thanks. If you search around on Qiagen's website, and go to the product pages for the various kits, you should be able to find documentation on how they work, and what the different buffers do.

I will and I might even ask Qiagen directly. For now, my follow up question if you are interested: http://biology.stackexchange.com/questions/40198/what-evidence-is-there-that-common-pcr-dna-purification-kits-can-isolate-dna-fro

I think the right answer is "they don't". If you were doing pcr on some sample that you didn't know much about, you would certainly include a positive control, meaning a primer that should amplify something that you think is definitely present in your sample. It you get a band for the positive control, then you know you successfully were able to lyse the cells and nothing else interfered with your pcr.
Most modern techniques on typical samples are quite good, so it's rarely a problem. But generally you would prefer to design your experiment to rely on positive results from pcr rather than negative ones.

The polymerase chain reaction is done on free DNA that has been released from the cell by a variety of chemical or physical methods.
There are no cells involved in PCR as the components you don't want are separated by high-speed centrifugation.
PCR also requires the cycling of heat as to aid in the unwinding of DNA for the binding of primers which also allows for the functionality of TAQ polymerase which is a high heat DNA polymerase isolated from the bacterium Thermus aquaticus.
So unless you're talking about a different reaction PCR does not involve any living organisms including cells. They are removed.

Thanks for your post! I have perhaps not phrased my question precisely enough. My question as mentioned in the edit can also be phrased: "can we be sure that the isolation techniques will not discard possible targets, e.g. from macrophages?" Taking your post the answer seems to be: "we can be sure that isolation techniques will discard most relevant targets." Can you elaborate?


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Why would the Camargue horses have a white coat while the foal have a dark coat?
The current issue of the magazine Lonely Planet India(Oct 2013) presents a photo of Camargue Horses which are white coated and mentions that they are born with dark hair. A search shows that the foal are either black, grey or brown coated and gradually turn white around the Third/fourth year.
What could be the possible advantage of that kind of difference between the young and the adult?

I will have to do more research into the Camargue, but many animals have pigment changes over time just because they haven't developed the melanocytes to have adult coloration.  It simply isn't advantageous to spend energy on coloration when you're growing bone. And sometimes the melanocytes are hormonally stimulated. A possible behavioral selection could be something like "foals are not seen as mate competition until their color changes," and thus avoid competitive behavior of say a large alpha male.

Our own hair will also turn gray (indeed, yours seems to be on its way, if you don't mind me saying so) but there need not be an advantage to that; if it doesn't affect our ability to have successful children it probably doesn't matter much to evolution or selection.  When it comes to animals that interact with humans, though, many traits are selected for, even those without an advantage, in fact often we select for disadvantageous traits (if they were advantageous, we might not need to select for them in the first place).
As this 2008 paper from Nature Genetics goes into, the "graying with age [phenotype] is an autosomal dominant trait" overwhelmingly caused by "a 4.6-kb duplication in intron 6 of STX17 (syntaxin-17) that constitutes a cis-acting regulatory mutation".  To answer your question, though:

The Gray horse provides a notable example of how humans have cherry-picked mutations with favorable phenotypic effects in domestic animals... The prestige of riding a white horse has thus led to selection of the Gray-causing mutation by humans; this mutation is by far the most common cause of white color in horse.

This phenotype is actually often detrimental, as white/gray horses can have eye problems and higher rates of skin cancer; Camargue's are particularly susceptible to melanomas, present in around 70% of horses by age 15.  There does seem to be some benefit, though, as the second paper I linked suggests these horses are subject to fewer fly attacks.
EDIT: I seem to have been unclear and you seem to have misunderstood my answer.  Let me break it down:

Camargue horses are gray.
Gray color is a simple mendelian autosomal dominant phenotype caused by a loss of pigment in adulthood.
Gray color is generally bad but sometimes good; either way, it's probably not a huge detriment.
Camargue horses are homozygous for gray color.
Humans have in the past selected for gray color in horses in general.

That is an example of why something negative or pointless might persist, as would a dominant inheritance pattern that is fixed in a population: negative traits that are only weakly deleterious have a non-negligible chance of fixation.  Asking what the advantage is is somewhat of a moot question because every single Camargue horse has the trait; asking what caused a seemingly-negative phenotype to become fixed is a good question, but likely an unanswerable one as it occurred long ago.
As a side note, how long a breed has been in an area does not simply imply breed purity.  Here's a quote from the International Museum of the Horse:

Through the centuries many armies have passed by the Camargue, including the Greeks, Romans and Arabs. The horses brought with these armies influenced the Camargue over time. It has even been suggested that the Camargue has had some influence on the early breeds in Spain as armies took them back home.

Your answer is not acceptable. 1. The Camargue Horses are neither 'cherry-picked' or cross bred. Read these quoted lines from Wikipedia "The Camargue horse is an ancient breed of horse indigenous to the Camargue area in southern France. Its origins remain relatively unknown, although it is generally considered one of the oldest breeds of horses in the world".

You are suggesting that the hair of the horse ages before the horse even before it reaches puberty?  There has to be a better explanation. It looks like you have cherrypicked the answers. Is it possible to get an answer from someone with some insight in this site?

@RamManoharM I expanded my answer, but I don't know where you find reason to accuse me of "cherry-picking" or find a mention of puberty.  I simply mentioned our own hair color as an example of seemingly-drastic changes that need not be selected for.

@RamManoharM  2 might not be to far off.  It's almost certainly related to the KIT gene (CD117, dealing with the SCF).  Early coloration being caused by in utero influences on the cells, but the later lack of coloration is cause by the failure of melanocyte migration.


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Does methanol fixation deform cultured cells?
I use methanol fixation (@ -20⁰C for 10 minutes) when performing immunofluorescence assays on cultured cells. Generally speaking, this results in very good antibody staining. However, the cells tend to look markedly different than they did under phase contrast microscopy prior to fixation. This has my PI worried about how these images will be received by reviewers.
How does a methanol fixative work, and why does it cause a change in cell morphology? Are there parameters that can be adjusted to mitigate this effect?

Give me one last bit of information, and I will throw up a re-write of why MtOH causes changes in cell morphology, why I really think you should be using acetone, and the unmasking information I already put up.  What are you culturing the cells on, and what are you fixing them on?  Is it a PP chamber slide, trans-well, what?  Direct trouble shooting a specific problem is a cornerstone of SE sites, check out stack overflow if you haven't looked at it already.  If you want a more general answer, be prepared to show what research you have already done to answer the question yourself.

Yes, fixation of almost any kind can have effects on morphology.  When you take a free flowing, protein spiked fat-blob (ie cell membrane), and make it rigid, you are going to get some differences.
A fun visualization of this can be done by wrapping cellophane/shrink wrap around a serological pipette and dipping it in a dry ice and methanol bath.  It's something I show students all the time before getting into histology work.
Back to what you need.  What cells are you using and what are you growing them on?  Have you tried acetone fixation and you know it destroyed your plastic?  The amount of time and concentration needed for acetone fixation is low enough that most modern chamber slides can handle it.  Do you need the membrane to be permeabilized in fixation?
I don't know the main effect you are measuring, but assuming you have good controls, you should be able to see it past the fixation.  The effect in your controls should be similar to your treated groups, unless the treatment is specifically effected by your fixative.
Moving to some more general recommendations.  I think you should reconsider the fixative’s you’ve ruled out, because I am unconfident about their application.  The base source I always recommend students to turn to is the Wiley "Current Protocols in X" book that is relevant to the task at hand.  In this case, I think the most relevant text is Current Protocols in Immunology, and the appropriate unit is 21.4 "Immunohistochemistry."
I think a particularly useful, and relevant to this question if you are concerned with formalin masking, is Table 21.4.3.

That is for human antigens, but the “antigen retrieval” process works for murine cells in most cases.  You are right that you antigen could be blocked by formalin, but the solution is often unmasking as MtOH are often more harsh to morphology.  To support protocol 1:

Formalin or paraformaldehyde fixation cross-links protein, often
  making antigens inaccessible to specific antibodies. The antigen
  retrieval protocol, which combines high temperature with buffers at
  different pH, has been very effective in unmasking antigenic
  determinants which may be masked in fixed tissue or fixed cell
  preparations, particularly determinants within the nucleus or
  cytoplasm (Taylor and Shi, 2013).

My modification of the protocol is:
Sodium citrate buffer: 0.1 M citric acid/0.1 M sodium citrate, pH 6.0

Wash sample on a liner rocker for 5 min w/2x culture volume of d.i. H2O
Place your slides in 1mM citrate buffer in Coplin jars with the lid slightly off
Weigh the jar in final state so you know how much water to add back
Microwave 4-7 min.  You don’t want it to go to a boil, and you need to remember to allow for venting
Weigh the jar again and add back water
Microwave another 4-7 min, again you don’t want a boil
Take out of the oven and allow it to come to room temp by sitting it on the bench.  If you do it too fast you’re going to disrupt the membrane.
After it’s back to room temp, wash 2x PBS+0.05% Tween20 for 10 min each
Try your IHC again, it will probably work now.

It's a surface antigen on endothelium, so I'm trying to leave the membrane as intact and possible. Acetone ruins our culture vessels pretty quickly. How long do you fix for? I am generally against the use of kits. They're too much of a black box (one of the many reasons I dislike the growing role and dubious business practices of some vendors). If you check the MSDS on that life technologies kit, you'll see that the fixation medium is just formaldehyde. Which I have already tried once, but am going to give another go tomorrow.

@Chastain First, you should almost never be using straight formaldehyde on an endothelium. You should be using a buffered formalin, about 10% in concentration. Second what's the tissue source? (Animal/organ)

I will also note that side by side with a buffered formalin and tissues from BALB/c mice, we saw much better preservation with the kit.  Could just be how they are buffering things.  I will post up the MtOH and Acetone protocols I use after my morning meetings.

Thank you. It's murine and isolated from several different organ systems. To be clear, the question is not about tissue section morphology. Morphology of sectioned and stained tissue is fine. This question is specific to cultured cells. Which is one of the reasons that it is so frustrating.

This perhaps could be settled out better over chat.  Histology, fixation, and IHC are things that have to be highly optimized. My staining of primary human lung epithelium was tweaked over several years and labs. I understand you are trying to fix cultured cells. Is it a monolayer or transwell system? From the sound of your last comment, you are co-culturing multiple types of cells (as they are from different tissues). Are they primary or imortalized? I'm trying to get you a solution rather than a general answer on fixation. If you don't want 1, I can write up an in-depth answer on fixation.

I'm new to stack exchange, so I'll have to figure out how to use the chat. To be clear though, and maybe I should edit my original question to reflect this, I am looking for the general principle. I love to chat about my work, but that seems outside the scope of the q & a format here. I want to know 1) if methanol fixation deforms a cell (I think that's been answered as a resounding yes!), 2) how methanol fixation works and why it deforms a cell, and 3) variables, if any, that enhance or mitigate this effect. It's important to me to understand the principle at work here.


text:
What sample sizes are ideal for carrying out Bayesian Skyline Plots?
I am interested in creating Bayesian Skyline Plots to look at demographic changes in certain population groups. However, these populations have very little within population variation. Around 5 polymorphic sites within each population and one dominating haplotype that comprises almost 90% of the population (the rest being rare haplotypes). Because, BSP uses Coalescent processes for the calculations, the sample sizes would have a effect on the results. I wanted to know if sample sizes of around 30 (for DNA alignments of 700bp) would be ideal for carrying out the analysis and expect reasonably good results ?

What software are you using?

Note that any plot is just a visualization method. You should be asking about sample sizes for reliable analysis. I am not an expert at this but according to wikipedia, Coalescence time and heterozygosity are dependent on population size. You have to clarify what you mean by "results".

I found a PLOS one article that might help. In this they used similar numbers you're planning to use (if i get your question and the article's data right).

The influence of the sampling scheme was investigated by drawing 40 samples in three different ways: 1) all 40 samples from a single deme, 2) 4 samples from each of 10 demes and 3) one sample from each of the 40 demes in the structured population. These correspond respectively to the local, pooled and scattered

(taken from the above linked article)
So depending on your sampling scenario your 30 samples seems to okay.
Also this is a link for the BEAST FAQ that might be helpful.


text:
Is H antigen considered as an agglutinogen?
A and B antigens which have the potential to cause agglutination in certain cases are called agglutinogens. But, as far as I know, H antigen cannot give rise to agglutination. So can it be said that H antigen also is an agglutinogen?

H antigen is a precursor for A and B antigens, and hh (recessive) individuals do not express this antigen. See Bombay blood group. This is an example of epistasis. Since H antigen is present in all A,B,AB and O blood groups, transfusion from any of these to an hh individual will cause a haemolytic reaction. Anti-H antibodies are agglutinins and have been referred to as agglutinins in the scientific literature.

The fairly strong anti-H agglutinins present in 'Bombay' and
  'para-Bombay' individuals (see 5.5.6) are generally accompanied by anti-A and anti-B, neither of which can be separated from the
  anti-H specific antibodies.

 Schenkel-Brunner, Helmut. Human blood groups: chemical and
  biochemical basis of antigen specificity. Springer Science &
  Business Media, 2000. e-Book ISBN: 978-3-7091-6294-1


text:
Gene and alleles
This is a multiple choice question:
Consider a gene, ABC, which codes for an enzyme involved in the metabolism of sugars. There are two known
alleles of this gene, ABC1 and ABC2. Which statement correctly describes the relationship between the ABC gene
and the ABC1 and ABC2 alleles?
a. The gene is a sequence of amino acids and the alleles are a very similar sequence of amino acids.
b. The gene is a trait and the alleles are a sequence of amino acids.
c. The gene is a trait and the alleles are a nucleotide sequence.
d. The gene is a nucleotide sequence and the alleles are a sequence of amino acids.
e. Both the gene and the alleles are a nucleotide sequence.
I thought the answer is b, but the correct answer is e. I can't figure out why so. Does anyone know?

Alleles are basically subtypes of a gene. At the time of Mendel, the molecular nature of inheritance was not known so the original definition of gene refers to "some" inheritable molecular entity inside the organism that is responsible for a trait. Alleles are different "flavours" of a given gene. For example there is a gene for flower colour, there can be different alleles which give rise to different colours (this is a highly simplified example). Genotype is a configuration of alleles whereas the phenotype is the effect that is seen.
With the knowledge of molecular genetics superimposed on these basic concepts a gene would basically be a well defined part of the genome (DNA) which is responsible for a molecular trait. Alleles are the actual sequence variants of this genomic region (not considering translocations here).
This is my justification on correctness and incorrectness of different points, based on current knowledge of molecular genetics:
a. Incorrect. Genes need not necessarily code for proteins. There are non-coding RNAs
b. Incorrect. Trait is a qualitative feature. Phenotype is the manifestation of a trait. Genes and genotypes are the causes of a trait and not traits themselves.
c. Incorrect. Same as above.
d. Incorrect. As per the definition, alleles are variants of a gene and they have to be of the same molecular nature as the genes. However, if we forget the semantics then this may seem like a more apt definition because the variations in traits arise not just because of the gene sequence but also the amino acids. However there is a flaw in this logic and the counter points be:

This scheme would not be inheritable (RNA editing/alternative splicing/PTM etc).
It has been shown that even synonymous mutations can have different phenotypes. (Plotkin and Kudla. 2011)
This does not consider non-coding RNAs

e. More or less correct (not in a very strict sense but fine for most discussions. Gene is not really a nucleotide sequence. Gene is an annotated region of a genome which has a defined sequence. This is similar to saying that object is mass. Object has a mass; object is not mass. This is just semantics and, as I said, not too important for general discussions.)

Both genes and alleles are sequences of DNA. A gene will code for a trait, say hair colour, while an allele will be the variants of that gene (say the alleles coding for blonde, brown, black, and red hair).
It's almost like cookbooks: two cookbooks (the DNA) might have a recipe (gene) for bread but they use slightly different instructions (alleles) resulting in different breads (traits).

That's a very good analogy imo


text:
Techniques to show a protein heterodimer-DNA interaction
It is known that c-Jun and fos dimerize to form AP1 factor that binds to a sequence on DNA containing PyPuGACGTCNNNNGAGGTCPyPU.  In esophageal cancer cell lines there is no expression of the fos gene. However we still see expression of genes regulated by AP1 factor.  One of the possibility is another isoform of fos is expressed in these cells (e.g. fra1).

How will you check that fra1 is expressed or not in these cells.
How will you check that the heterodimer bound to the given DNA
contains fra-1 along with c-jun.

My thoughts on the above questions
For first question we can do immunofluorescence assay or we can do DNA affinity assay in which we chemically synthesize DNA sequence and bind it on agarose gel column pass through it protein extract of oesophageal cancer cells and our fra-1 protein if being expressed in cell line will bind to DNA. later we can elute protein DNA complex and by anti-fra1 antibody we can check if it is being expressed. But the drawback of both these technique is we should have knowledge of epitope of fra-1.
For second question - we can perform ChIP using antibody tagged Against c-jun and then eluting out DNA bound to c-jun. Then we can simultaneously label the complex with anti fra-1 in one sample and anti fos in another sample. We can check for results. If fluorescence is obtained in case of fra-1 sample that means both c-jun and fra are bound to probe and not fos.

You should add your thoughts onto the post directly and use the homework tag if it is a homework. Homework questions are on-topic as long as the OP is showing effort for answering the question himself/herself

I'm going to throw RT-qPCR and EMSA "supershift" out there, but there are other methods, with their own pros and cons.

How will you check that fra1 is expressed or not in these cells.

RT-PCR is a quick and an easy method to check for expression. However, it only tells you about the RNA level expression and you would have to work with the assumption that RNA concentration is directly proportional to protein concentration.
Considering that you do not have an antibody for fra1, you can go for mass-spectrometric protein quantification. It is good for a high throughput analysis and is generally wasteful when you just want to study one protein. However, people do carry out targeted proteomics to study a few proteins.
Alternatively, with some effort you can actually develop an antibody for fra1.

How will you check that the heterodimer bound to the given DNA
  contains fra-1 along with c-jun.

Showing ternary complexes (fra-jun-DNA) are a bit difficult. It is relatively simple to show pairs. You can show that two proteins form a complex in vivo by doing a co-immunoprecipitation followed by detection by either western blot or mass-spectrometry.
You can also show interaction by biophysical techniques involving FRET and SPR but these cannot be performed in vivo (at least SPR. FRET can be done but there are limitations).
You can show the DNA-protein interaction by ChIP-seq as you mentioned in your question.
Showing all the three together is a bit tricky.
You can do it using in vitro assays that employ fluorescently labelled (one GFP and other RFP; or FRET pairs)/custom epitope tagged (for e.g. one FLAG and the other HA -tagged) proteins and a known DNA sequence generated by PCR/DNA synthesis. You can use EMSA along with antibody/fluorescence based detection to show the ternary complex.
You can indirectly infer the presence of the ternary complex when the ChIP-seq using antibodies for both the proteins shows up the same DNA sequence in the reads. You can also express a FLAG-tagged (or any other tag) protein, if in case you do not have an antibody for the proteins.

Nice list of many techniques, I'd just like to mention that you could also label the the DNA with biotin, an isotope or something else to suit your needs.

Can you clarify on the term "ternary", also ditto ^^

@hello_there_andy "ternary" means a complex of three components.


text:
Do ants really dispose of their own dead, and how/why?
I just read this from Wikipedia's Swarm Behavior article:

Despite the lack of centralized decision making, ant colonies exhibit complex behaviour and have even been able to demonstrate the ability to solve geometric problems. For example, colonies routinely find the maximum distance from all colony entrances to dispose of dead bodies.

There was no citation.
Is it true that ants dispose of their own dead? If so, how and why? I would've though it would be much better to just eat their own dead, which would be a "free" food source allowing the colony to reclaim some protein and nutrients. It would cost much more energy (and risk) to transport dead ant carcasses away from the hive.

If you want to watch something scary, google for "youtube cordyceps" for a clip from "Planet Earth".

Eating the dead makes far more likely you will catch whatever killed them. They transport and dispose of the dead for the same reasons we do, to reduce disease vectors. They dispose of their garbage for much the same reason, in fact they often dispose of both in the same place even if they separate the two in distinct piles. Hygiene is very important is colonial insects because you are constantly surrounded by other ants which makes disease transfer very easy. Fungal infections are particularly problematic. They also have specialized toilet sites for the same reason.
Note only some ants transport the dead outside, other put them is special chambers far from normal traffic and reserved for that purpose and refuse disposal. In either case they are moving them away from casual contact.
Source
Source 2

Ants do of course have emotions, emotions are the basic way the brain functions, but no they do not "remember the dead". humans bury their dead because being around the dead make us uncomfortable, our brains evolved to make us feel uncomfortable around corpses because they are disease vectors. So yes it is ultimately for the same reason.

pleasure and aversion are the basic building blocks they underlie all learning, emotions exist in any creature capable of learning. as for the human corpse, I am unsure what you mean by "it's most an emotional"  but corpse aversion it is the explanation for why the uncanny valley  a well documented effect http://www.sciencedirect.com/science/article/pii/S0747563210001536

You can also consider things like the disgust/revulsion reflex triggered by decaying corpses, which is as universal https://www.ncbi.nlm.nih.gov/pubmed/17630362 it is not hard to see how it would evolve as part of our behavioral immune system. Feel free to open a discussion to explore this further.

I'd like to add a +1 to John's comment re: animal emotions. Charles, you just specified that a physiological response while panicking was not an emotion. What do you think panic is?


text:
Is there a reliable source for storage and stability of reducing agents like DTT?
Reading the literature on DTT, one is confronted with a confusing mass of papers; some claim that a 1M solution in water is stable, other papers say it is not. I use the reaction with DTNB to show that the DTT is still good, but that is cumbersome.
Is there some source of reliable (as in data driven) information on how to store compounds like DTT and other reducing agents?

Thiol-Based Reducing Agents
The instability of thiol-based reducing agents in solution is due to their propensity to form disulfide bonds by the following half reaction:
$$\ce{2RSH -> RSSR + 2H+ + 2e-}$$
It is clear that disulfide formation requires:

Deprotonation of the thiols to form thiolates (RS-).
An electron acceptor.

Therefore, solutions of thiol-based reducing agents would be most stable when:

The pH is buffered to where the thiol form predominates. The pKa of the thiol in β-mercaptoethanol is 9.5 and in dithiothreitol are 9.2 and 10.1  (Whitesides et al., 1977). Buffering the pH lower than this should increase stability.
Electron acceptors are removed. In aqueous solution, the electron acceptor is molecular oxygen. Degassing the buffer and storing the solutions under an inert gas such as argon should increase stability.
Reaction catalysts are removed. Divalent metal cations, especially copper (II) (Cu2+), catalyze oxygen dependent disulfide formation, even in trace amounts (Smith et al., 1994). Therefore a chelator such as EDTA should increase stability.
Temperature is low. As with most chemical reactions, rate increases with temperature.

The stabilities of the thiol-based reducing agents β-mercaptoethanol (β-ME), dithiothreitol (DTT) and glutathione (GSH) have been determined in solution as a function of pH at 20°C (Stevens et al., 1983):

...and temperature at pH 8.5:

...as well as in the presence of copper (II) and EDTA at pH 8.5 and 20°C:

Below is the tabulated raw data (this is the actual data from the paper which I used to create the above graphs, but take note that I found it difficult to plot “>100”):
                               Half-life (h)
pH    Temp (°C)    Additive    β-ME    DTT    GSH
6.5   20           -           >100    40     16
7.5   20           -           10      10     9
8.5   20           -           4.0     1.4    1.3
8.5   0            -           21      11     8
8.5   40           -           1.0     0.2    0.2
8.5   20           Cu(II)      0.6     0.6    1.2
8.5   20           EDTA        >100    4      70

As expected, stability decreased with increasing pH and temperature, and also with the addition of copper. Stability increased with the addition of EDTA.

Non-Thiol-Based Reducing Agents
Han and Han (1994) studied the effect of pH and buffer composition on the solution stability of tris(2-carboxyethyl)phosphine (TCEP). Though this is not a thiol-based reducing agent, it is still subject to oxidation. However, it is quite stable in solution:

pH    Solute/Buffer     % TCEP Oxidized After 3 Weeks
7.5   Tris-HCl          18.7 ± 0.6
8.5   Tris-HCl          16.8 ± 0.5
9.5   Tris-HCl          14.5 ± 0.5
8.2   borate             6.6 ± 0.2
10.2  borate             5.3 ± 0.1
6.8   HEPES             14.8 ± 0.4
8.2   HEPES             13.6 ± 0.3
9.7   CAPS               9.8 ± 0.3
11.1  CAPS               4.3 ± 0.1

Stability increased with increasing pH, though the effect was not large. Interestingly, the buffer composition had a larger effect on stability which was especially pronounced with phosphate:

        % TCEP Oxidized After 72 Hours
 pH     0.15 M phosphate    0.35 M phosphate
 6.0    not determined      11.2 ± 0.3
 6.8     8.1 ± 0.4          56.8 ± 2.3
 7.0    23.5 ± 0.9           100
 7.2    33.4 ± 1.0           100
 7.4    56.5 ± 1.4           100
 7.6    56.8 ± 2.1           100
 7.8    48.9 ± 2.5           100
 8.0    36.6 ± 1.7           100
10.6    not determined      11.7 ± 0.5
11.6    not determined       9.1 ± 0.4

References
Han JC, Han GY. 1994. A Procedure for Quantitative Determination of Tris(2-Carboxyethyl)phosphine, an Odorless Reducing Agent More Stable and Effective Than Dithiothreitol. Anal Biochem 220(1):5-10.
Smith RC, Reed VD, Hill WE. 1994. Oxidation Of Thiols By Copper(II). Phosphorus Sulfur Silicon Relat Elem 90(1-4):147-154.
Stevens R, Stevens L, Price NC. 1983. The stabilities of various thiol compounds used in protein purifications. Biochemical Education 11(2):70.
Whitesides GM, Lilburn JE, Szajewski RP. 1977. Rates of thiol-disulfide interchange reactions between mono- and dithiols and Ellman's reagent. J Org Chem 42(2):332-338.

Very useful references, thank you. Do you know if there is a similar, probably more recent article that also addresses the stability of TCEP?

@Guillaume I might have some; I’ll check when I’m at work tomorrow. I work with TCEP frequently and, in my experience, it is very stable in aqueous solution, though I don’t have any quantitative data.

@Guillaume I've updated the answer to include the data about BME and glutathione as well as TCEP.


text:
Problems with SDS-PAGE, no separation - what could be the problem?
I work in a lab were we have a common stock of 30% Acrylamide, TEMED, TRIS-HCl (pH6.8 and pH 8.8.) and a 10% (w/v) SDS solution pH 7.2.
We use the recipe for loading buffer (which we of course mix with BME before use):

3.15 mL MilliQ water
1.25 mL 0.5M Tris-HCl
2.9 mL 85% Glycerol
2 mL 10% SDS
0.4 mL 0.5% (w/v) bromophenol blue

Our problem is that there is no separation.
I have tried making 2 gels because I though I might have mixed the separating and the stacking gel, but I made sure i did not do this the second time. Then I asked someone else to make the gel, and it did not work for them either (nothing on the gel, no samples no ladder).
Then I remade the 10% SDS solution and both the TRIS buffers, but again something strange is going on as we get nothing on the gel.
The gels are obviously polymerized as its a solid gel when you take out to stain and destain, and I dont understand how the loading buffer can do this even if you mess it up (because you should see something on the gel as long as your sample in the well, even without denaturing). We obviously remade the loading dye and tried again, but this does not work.
I know there is protein in the samples, as I am putting cell pellet and purified protein (which I have tested the catalytic activity of) on the same gel.
Does anyone have a rational explanation for this, or suggestions of what can typically be wrong.
Bellow is a picture of the gel as it looks when its almost done running at 200V (80mAmp) for approx 40min. When we stain and destain there is nothing on the gel. Also I think the color (bromophenol blue) looks as if its running a bit strange on the gel, not as it normally should, see picture below, but perhaps I'm getting a bit paranoid.

Also try using a pre-stained protein ladder in one of the wells. (Preferably one that comes ready-mixed and doesn't need loading dye/heating/BME.) That should separate issues with you gel/running conditions from any issues with your protein/loading dye/staining process.

Are other people in your lab able to see protein in their gels? Not your specific protein, but any other protein that they're working with. In case you don't have pre-stained markers, this should also tell you whether it is a gel/staining issue or a problem with your specific protein. Also, how much protein are you loading?

Thanks fot the input, after readjusting the pH of everything and making new running buffer with new SDS solution the separation seems to be back. Its still a bit of a mystery, but perhaps the SDS (suddenly) got old and since its in both the loading dye, running buffer and gel - it just did not work. I have been reading a bit and it seems that some say that the 10% SDS should not be older than 6moths, ours was 5 years old but had been working just fine up till one week ago.

After readjusting the pH of everything and making new running buffer with new SDS solution the separation seems to be back.


text:
Hamming distance between two DNA strings
By definition from Wikipedia, the Hamming distance between two strings of equal length is the number of positions at which the corresponding symbols are different. In other words, it is the number of substitutions required to transform one string into another. Given two strings of equal length, compute the Hamming distance.
So to find the how many differences there are between two strings, say we have two DNA strings A and B:
A = TGACCCGTTATGCTCGAGTTCGGTCAGAGCGTCATTGCGAGTAGTCGTTTGCTTTCTCAAACTCC
B = GAGCGATTAAGCGTGACAGCCCCAGGGAACCCACAAAACGTGATCGCAGTCCATCCGATCATACA

Throw them to two arrays A and B. Therefore, If
(= count 0)
(cond
   [(symbol=? A[1] B[1]) /do nothing just move on to the next pair of string]
   [else (add1 count)] ;; counter:= counter + 1;

output would be the counter.
Is this the right logic flow to go with this problem? Any suggestion to write this on python or Scheme?

I think for the programming side of things, this question should be asked (at least in part) in http://stackoverflow.com

This question does fit into bioinformatics, which is on-topic here. That said, it really has nothing to do with DNA and you might get a better response at stackoverlow.

By the way, this sounds a lot like a homework question and there is a tag for that here.

Your logic on the problem seems correct. The wikipedia article about the Hamming distance describes both a python and a C implementations. The python version assumes two strings of equal length:
def hamming_distance(s1, s2):
    #Return the Hamming distance between equal-length sequences
    if len(s1) != len(s2):
        raise ValueError("Undefined for sequences of unequal length")
    return sum(ch1 != ch2 for ch1, ch2 in zip(s1, s2))


text:
Skin color explanation?
I do not understand how skin color in humans work, even after taking a basic genetics course and reading some on wikipedia.
From what I understand, skin is color based on multiple genes that control melanin and other products that affect appearance of skin. So then their are multiple alleles for "skin color" and also some of these alleles can be varying in their expression/inheritance? based on the other alleles presence?(epistasis)
So then a black person and a white person could have various skin colors (phenotypes?) And the offspring of these children could then have varying offspring as well?
Diagrams would help me maybe in explaining the concept.

I'm still a little uncertain as to where your question lies.  Are you wondering how different melanin types and levels create different skin phenotypes (of any animal, no real reason to limit this to humans), or are you interested in how genetic inheritance effects melanocytes?

I have the same problem. Do you look more into melanin production and how this is regulated or more into the genetic reasons for different skin color?

@AtlLED           Ummm, I want to know the underlying genetic components(how the genes work) more than their product(cells, proteins, etc..)

This is relatively complicated.

@RoSiv So is your question "How do genes work?"

The genetics of pigmentation is relatively complicated, as the pathway for the pigmentation (regulation of the pigment production, ratio between the melanins, maturation, trafficking and distribution of the melanins from the melanocytes to other cells) is quite long and also subject to different regulations. All the mutations found to date (at least to my knowledge) affect single parts of this long process (so for example MC1R mutations are affecting whether eumelanin is made, while Rab7 and Rab27a is most likely involved in the distribution of the melanosomes). Not all gene functions in this network have been understood so far.
Population studies have shown that the three main ethnic subgroups (west-africans, asians and europeans) have developed independently from a common ancestor with different (and independent) mutations in each arm. See the figure from reference 1 as an illustration, it also shows the genes involved in each arm:

Pigmentation is a protection against UV light, so there is a strong selection against mutations in areas with high UV light (west africa for example). The general theory is that in areas with less UV light the skin got lighter to allow the production of vitamin D (for which you need UV light). If you want to go through the single genes and their influence on pigmentation, please have a look at this Wikipedia page, which gives a good explanation. Also have a look at the references.
References:

Unpacking Human Evolution to Find the Genetic Determinants of Human
Skin Pigmentation
The genetics of pigmentation: from fancy genes to complex traits.
Molecular genetics of human pigmentation diversity
Human pigmentation genetics: the difference is only skin deep
The etiology and molecular genetics of human pigmentation disorders

This HHMI Biointeractive Video gives one of the better explanations I have seen on the genetics of Human Skin color. The Biology of Skin Color It is actually the destruction of folate, which is known to lead to birth defects and sperm fitness that selects for Eumelanin in regions with high levels of UV.

Ok, several genes are involved, which increases the combinatory. But still, this should give a finite number of genotypes, and thus of phenotypes, right ? -> is the combinatory just large enough (estimation?) so that we have the false impression of continuous shade ? Or is there something else occurring, making the shades really continuous ?


text:
Lizards are sometimes called cold-blooded because they depend on their surrounding temperatures to keep them warm.
Predict how the composition of the cell membrane of a lizard might change from winter to summer. Justify predictions.
Can someone please help me figure this out. Does it have to do with fluidity?

I didn't really get your question. Is it: "Cell membrane composition (or other properties of the membranes) depends on temperature. Why is it so?"

Be sure to tag homework questions with the homework tag.  It also helps if you let us know what level class you are taking.  To your question, look up how cell walls in general help maintain homeostasis.  Edit your question with what you find, and feel free to answer your own question if you figure it out.  Give it a little more effort and come back to us with what you found.

Just going to follow up on your thought here.
The fluidity of the cell membrane does vary with composition.  Off the top of my head there are four main components of the animal cell membrane:
1) proteins.  membrane proteins are responsible for the import and export of nutrients, waste, information etc from the cell, so lets assume that this needs to be fairly constant part of the composition while the seasons change.
2) cholesterol and similar compounds.  These are flat an inflexible compounds which will tend to lower the fluidity of the membrane.
3) unsaturated fatty acids and lipids.  These are long carbon chained compounds with some double bonds in them.  Because the double bonds cause the chains to be less mobile, but they don't pack well as they are not as flexible, so tend to make the membrane a bit more fluid.  Think about how butter (mostly saturated fats) is solid at room temperature while olive oil (monounsaturated oil) is liquid.
4) saturated fatty acids and lipids.  These are completely mobile carbon chains.  Without any double bonds, these tend to be the more flexible and can create a more tightly packed membrane, resulting in a stiffer membraine.
You can imagine for the cell to maintain the same flexibility at the lower winter temperatures (4) and (2) would tend to decrease and (3) might tend to decrease.
Does this make sense?


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Plumage as indicator of health in birds
Anybody know which traits of bird plumage are accurate indicators of a birds health? For example feather coloration, feather brightness, amount of feather wear, number of feathers, moult timing, moult duration etc.

This resource provides a short list of which general components of plumage are good indicators of a healthy bird.

Your pet's plumage should be. . .
Soft - feathers should be strong, yet flexible.
Smooth - no rough feather shafts; no uneven or split edges
Glossy - sheen extends over the entire coat
Full - thick, where it needs to be
Vivid - color should be deep, bright and uniform
Clean - free of dust, dander, waste, or soil
Parasite-Free - no lice or mites

Also, this website page discusses the signs of malnutrition or disease in plumage in more detail.
Stress bars, dark lines which transect a feather, are directly caused by an inefficient absorption or lack of required nutrients. Stress bars can be found on all feathers during the period of malnutrition, and remain even after this period.
Assymetrical moulting is also a sign of bad health. There should not be any bald patches present during the moult.
The resource states that if a moult is delayed due to malnutrition, an incorrect photoperiod or a metabolic problem, plumage will become depigmented. However, the appearance of "dirty" feathers can also be due to overpreening or damage done to feathers by physical contact.
Finally, a change in the usual plumage color of the bird in question is indicative of a lack  or oversupply of a particular nutrient.


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Why are urban birds still scared of humans?
I understand that animals are naturally scared of humans because we basically slaughtered every living thing we found in our way for millennia. How long does it take for them to genetically 'forget' we are a threat?

What's your evidence that an individual animal who has never encountered humans before is "scared" of humans more than other species?

@MCM I am not really sure I understand? I was referring to animals that have encountered humans before. I am mostly curious about the recent phenomenon of urban birds who have co-existed alongside with humans for a few decades now, and that for the most part these humans presented no threat to them. It seems to me that even after years of co-existence they still prefer keeping a safe distance from us humans.

@IsopycnalOscillation, if they have seen humans before then their natural fear response is justified.  In the city I live, I've noticed that some children are particularly cruel to birds constantly chasing them away.

a) Plenty of birds in the city will get very close to humans, relative to countryside birds (if you've ever been to Trafalgar square in London you will know how close birds are willing to get, try getting that close to a pheasant in a field!). b) It may be that "scaredness" (let's go with timidness) is not a genetic trait - or has a very large environmental component (i.e. largely determined by previous experience of human).

Perhaps counter-intuitively, fear of humans is often a learned behaviour. There are accounts, such as one made by Charles Darwin following his visit to the Galapagos islands;

In Charles Island, which had then been colonized about six years, I saw a boy sitting by a well with a switch in his hand, with which he killed the doves and finches as they came to drink. He had already procured a little heap of them for his dinner, and he said that he had constantly been in the habit of waiting by this well for the same purpose. It would appear that the birds of this archipelago, not having as yet learnt that man is a more dangerous animal than the tortoise or the Amblyrhynchus, disregard him, in the same manner as in England shy birds, such as magpies, disregard the cows and horses grazing in our fields.
From "The Voyage of the Beagle" - http://www.ling.upenn.edu/courses/hum100/Beagle17.html

Birds born and bred in proximity to humans will have learned that we are actually to be avoided.
Ducks and swans, for instance, (and this is quite anecdotal) live in very close proximity to a lot of humans in cities, and feel confident walking picking up food. However if you try to get close to one they are very quick to get out of the way. The only exceptions to this are learned behaviours, such as when someone stands and feeds them.
In the wild animals may be cautious of humans if we are much bigger than them, but if they have no experience at all of human contact they  may just ignore them. This quite often happens with film crews in Antarctica, as in the below image from the BBC's Frozen Planet series.

I guess I am biased because I would never hurt an animal. There may as well be other people out there doing just that - therefore perpetuating their fear of us. It's just so sad, when I go to the local creek, I would love to see more birds, I know they are there because I can hear them...

@IsopycnalOscillation I would guess that this is much beyond animal cruelty and probably goes as deep as innate behaviour. We hunted ducks and swans for food, ones that were innately fearful of humans survived and propagated and the next generation were predisposed to fear humans a bit more. Rinse and repeat for thousands of years. This would only be compounded by learned behaviour.

@GoodGravy Great point.


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What are chromosome dimers and how do they relate to recombination?
I am reading the book Molecular Genetics of Bacteria 4th ed. and I don't fully understand the concept/visual representation of dimer chromosomes. Consider the following quote (pg 35):

an odd number of crossovers occurring between any 2 sequences on the 2 daughter DNAs in the dimer will resolve the dimer, but an even number of crossovers will recreate a dimer.

In panel 2, did another recombination event occur to yield 2 daughter chromosomes with the same sequences? The colors made me think they each might not have had their original complete sequence.
Thank you for the answer to my crossover occurrence question and their relation to dimer formation and resolution.

Welcome to Biology.SE. I wanted to improve the format of your post but I was not exactly sure what you were asking. I've made an edit feel free to roll back if you don't like it. Also, if you accept the edit, you should definitely edit the <insert something> elements. +1 btw

The quote is not so specific to bacterial genetics as much as to genetic recombination in general. Have a look at this answer to understand why the question of whether an even or an odd number of recombination events occur between two loci matter.

Your question is rather broad. It would be better to ask about something specific that is confusing you about chromosome dimers.

Is your question why an even number of crossovers will recreate the dimer during resolution? That can be answered, though it would help if you explained why this doesn't make sense to you.


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If you measure the O.D. of cells at one wavelength, is it possible to find its equivalent O.D. at another wavelength?
For example, I measured the optical density (O.D.) of some cells at wavelength 550 nm but I should have used 600 nm. Is there a way of converting my O.D. to its equivalent in another wavelength?

It would be better if you explain what's OD?

It would be useful if you could tell us what you were using your measurements of optical density for?

O.D. can mean 30 different things, many of them being related to science and to optics (see here)! I'll assume you mean absorbance (for Optical Density I'd guess) or light scattering as @canadianer said
Each substance has its own absorption spectrum and reflectance spectrum. Knowing only the absorption is a given wavelength does not tell much about the absorption at another wavelength (at least not without some other a priori knowledge about the substance studied).
So, the answer is 'no'.

Your answer is essentially correct except that OD600 readings of cell cultures are a measure primarily of light scattering, not absorbance.

As already said by Remi.b, there is no way you can directly convert the O.D.550 to an O.D.600.
However, in case you already made numerous experiments using O.D.550, you may not have to do them all over again. You can try to draw a calibration curve of O.D.600 as a function of O.D.550:

take a cell culture (of the same cells you used for your experiments) and prepare several dilutions of it, so that their respective O.D.550 range from 0.1 to 1.
then, measure O.D.600 for each one of these dilutions.
then, draw O.D.600 as a function of O.D.550.

If, and only if, the points form a straight line, you can apply a linear regression to your data. You can then use this linear regression to convert your O.D.550 values to O.D.600 values. However, as pointed out be Jack Aidley in comment, it can be expected that the relationship is not linear. Provided that the curve is smooth enough, you can still calibrate to a curve.
Be careful:

this kind of conversion is totally empirical and will only apply to cell cultures of the same species in the same conditions.
this conversion only applies for values within the measured range of O.Ds.

Depending on what kind of experiment you were doing, it can be better to try this, and even replicate step one and two, or it can be better to start your experiments from scratch... Only you can tell

600 is traditionally used because it gives a more linear relationship between OD and cell count. I would expect that this would give a curve rather than a linear relationship; however, providing the curve is smooth and the conversions occurs within the measured range I do not thing calibrating to a curve is a problem

@JackAidley Thank you for the feedback. I've edited my answer accordingly.


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To what extent do gonads affect differences in gendered behaviour?
I'm not sure whether this question makes sense, but someone said that they result in gender differentiation on an organisational level in the brain. I'm wondering how important these changes are, in terms of magnitude of average differences in neural structure between males and females?

Can you specify whether you refer to humans? Because in different species (e.g. lizards vs mosquitoes) the importance of the gonads on behaviour changes  quite dramatically.

@havefun I was referring to humans, but would you be able to explain the other factors involved for lizards - and how the gonadal hormones play a role?

Yes
During embryonic life,there is formation of new synapses by a process termed as exuberant synaptogenesis.
Their differential formation in the males and females could be due to the effect produced by the gene PCDH11X/Y located in the allosomes.

PCDH11Y is a gene unique to human males that encodes Protocadherin 11Y, aprotein that guides the development of nerve cells. PCDH11X, located on the X chromosome, is common, in both sexes, to humans and our nearest relative, thechimpanzee; however, PCDH11Y, located on the Y chromosome, is unique to males.[3]
  ...PCDH11X/Y are cadherin family genes. They make proteins, involved in signalling, that attach to the surface of nerve cells.[4] PCDH11X and PCDH11Y, respond in different ways to Retinoic acid, a chemical involved in the development of embryos. The acid stimulates the activity of PCDH11Y but suppresses PCDH11X. This is likely one of the explanations for the differences between the brains of men and women.[3]
  [source]

About the parts of the brain which are deemed"sexually dimorphised" a few of them are:
1.hemispherical differences due to which inter hemispherical functioning is better is females
 whereas intra hemispherical functioning in males
2.hippocampal and amygdaloid laterisation

on average males had larger grey matter volume in bilateral amygdalae,hippocampi, anterior parahippocampal gyri, posterior cingulate gyri, precuneus,putamen and temporal poles, areas in the left posterior and anterior cingulate gyri
  ...females on average had larger grey matter volume at the rightfrontal pole, inferior and middle frontal gyri, pars triangularis, planum temporale/parietal operculum, anterior cingulate gyrus, insular cortex, and Heschl's gyrus; bilateral thalami andprecuneus; the left parahippocampal gyrus and lateral occipital cortex
  Source:Neuroscience of sex differences

Furthur,since synaptogenesis may occur throughout one's lifespan perhaps not as greatly as during embryonic life and as receptors for gonadal hormones are present in the brain it can be said that the gonads may affect the phenomenon,though mainly after puberty when their concentrations are higher than during childhood.

Regarding the third Wikipedia link, it would be best if you could include some significant passages. Links die, or the content might be removed, and it really is best to have relevant content available in the answer itself. You don't have to go overboard, but making the answer a bit lengthy isn't a problem if it's a good answer.

@Harris Weinstein thank u for ur suggestion..i will brief them.hope it helps.


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What is this web-like organic matter?
I saw this on my window couple of days ago. Does anyone know what it is? I have tried google image search with no luck.

scale would help a lot. I could be looking at an arctic oil spill satellite image.

Hi @Charles I would say it appeared out of no where I was out in the front lawn and noticed this it would have definitely caught my eye if it was developing and in its early stages

@John, I cropped the image but this is on my front bay window (outside) it is roughly 2 to 3 inches in length.


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Can methionine peptidase liberate selenomethionine?
Can methionine peptidase liberate selenomethionine from proteins? I'd be happy to know if any can that are active in humans, but am also especially curious about METAP2.

methionine peptidase is not specific enough. There are many different enzymes (from different organisms) belonging to that "family". Which one are you interested in?

I'd be happy to know if any can that are active in humans, but am also especially curious about METAP2.

Are you aware that METAP2 cleaves only N-term Met, and only followed by a subset of the standard amino acids?


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Identification of fuzzy insect in Pennsylvania
I live in south eastern Pennsylvania, USA. About a month ago I came across this very peculiar insect. Its body was about an inch long, it was fuzzy, and it had a long proboscis(?) and long orange legs.  I've been wondering about it ever since. Can anyone identify it?

Related: What kind of red and black insect is in the photo?


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Lowering annealing temperature in PCR
In the book "Lewin's Genes" there is the following quote:

Lowering the Tm of a PCR reaction—in effect, relaxing the reaction stringency and allowing primers to anneal to not quite
perfect hybridization partners — has useful applications, as well, such as in searching a sample for an unknown sequence suspected to be similar to a known one.

Why is this so? That is, why does lowering the annealing temperature in a PCR reaction reduce a specificity of hybridization, more precisely?

https://www.thermofisher.com/us/en/home/life-science/cloning/cloning-learning-center/invitrogen-school-of-molecular-biology/pcr-education/pcr-reagents-enzymes/pcr-cycling-considerations.html, see section "Primer annealing optimization".

Perhaps more specifically, see section "Effect of annealing temperature on primer mismatch" here: https://academic.oup.com/femsec/article/60/2/341/584515

Welcome to Biology.SE. The Biology.SE community has agreed that questions that show little or no prior research effort are off-topic on this site as are "homework" questions unless you have shown your attempt at an answer. Please [edit] your question and tell us where you've looked for answers, what you do know about the topic, and where exactly you still have questions. Under researched questions may be subject to down-voting and closure. Please take the [tour] and consult the [help] starting with [ask] for details.


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Qpcr negative efficiency
Hello Stack Exchange Biology world,
I have struggled with my qPRC results for over 5 months. I have a bunch of DNA samples, and I want to find the quantity of Pseudomonas syringae contained. My issue was the extremely high efficiency (sometimes even 1200%) after the run. So, I decided to make everything from scratch. I used a new set of primers and a new qPCR kit (kapa sybr fast qpcr master mix (2x)), and I switched the sterilized double-distilled water from the water filters in the lab to an ampoule one (to eliminate any contamination).
I incubated the bacteria again. I measured the OD=0.998A, before extracting this DNA (via NucleoSpin Soil, Mini kit for DNA from soil) with the following quality as 193μg/mL, 260/230 = 2,41 & 260/280=1,89, which seems high quality.
For the standard curve, I did dilutions of 1:10, 1:100, 1:1000, and 1:10000 using the proportion of 5μl of DNA with 45μl SDW from the ampoule.
So now I have everything new and a "high quality of DNA" to work with. Yesterday, I attempted to perform a qPCR (SYBR green), and the efficiency was -100%. What should be wrong at this time? Is it something that I can change in my 20μl reaction?
SD Water from ampoule 7μl
QPCR Master Mix 10μl
Primer F 0,3 μl
Primer R 0,3 μl
ROX high 0,4 μl
DNA 2μl

The machine I performed the reactions on is the Thermo QuantStudio 5 Real-Time PCR. Below, I am providing the experimental details as well as the qPCR results.
Thanks in advance for any suggestions to help me fix this issue.

Normally, if your qPCR efficiency was negative 100% I would say your dilutions were made properly just entered into the software in reverse order. But your amplification curves are pretty funky. How are you setting the threshold? And what is the baseline determination?  What about the melt curves? The protocol is strange too, in that it's taking multiple fluorescence readings per cycle. I wonder if that's screwing with the baseline readings. Typically just one measurement is taken after annealing/extension. Have you tried a 2-step protocol (deleting the 79C step)?

@MikeyC Thank you for your response. As I'm learning qPCR, I combined the KAPA SYBR® FAST Master Mix with a P. syringae protocol I found.  https://journals.asm.org/doi/10.1128/aem. The main issue was that the protocol used a different qPCR mix and annealing at 54°C compared to KAPA SYBR®(annealing at 60°C). I adjusted accordingly, but perhaps incorrectly. About the threshold and baseline determination, I used default settings. For my next run, I will only keep step 2 for fluorescence reading as recommended. Any further suggestions for refining my experimental method will be highly appreciated.

Run some of the qPCR results on a gel or have a look at the melt curves - you might well have more than one product being generated. Your std curve should have a little over 3 cycles between each point (copy number doubles each cycle so 3 cycles is 8x more copies to reach Ct) and the replicates should be quite tight. How long is the PCR product? It shouldn't need extension if it is <100 bp.

@bob1 I did, and you are absolutely right. Indeed, I have two zones on the curve amplicon. How do you know that? Wow, I am just surprised. Does it mean that my primers were not extended correctly? The primer sizes are 194 and 162 bp.

@KGee It was a suspicion (MikeyC noted it too), experience is a great tool... Your primers are 194 bp and 162 bp each? Or are those the amplicon sizes? Primers should only need to be about 18-25 bases for a highly specific reaction. The manual for the Kapa pol says extension of products less than 400 bp should need 1 second.

@bob1 Oh, I didn't get that (MikeyC) mentioned double zones. Obviously, you are an expert user. :) Do you recommend adjusting step 3 from 3 to 1? Will this extension fix the issue of the double zones? The f primer is 194 (22 bases), the R one is 162bp (20 bases), and the amplicon size is about 150bp. Thank you again!

Ah, I see - the primers are called "194" and "162". The sizes 20 and 22 are fine. You should go and optimize the binding of the primers in the Kapa mix; it might well be different to in another system.  A 3-step is fine. The Kapa manual says  For 3-step cycling protocols, anneal at optimal annealing temperature
for 20 sec followed by the minimum time required for data acquisition
at 72ºC according to instrument guidelines)  So you should acquire only at extension and for the minimum capable on your instrument. You'd need to check the instrument manual for this information.

@bob1 Oh, I see. Thank you once again for the response. Let's see. Then I hope to "get rid" of double zones this time  :P!!!


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What is the latest consensus on the causes of sexuality?
In 2019, scientists concluded that there is no gay gene. Epigentic markers of homosexuality have been disproven. The Exotic becomes Erotic Theory is an ancient decrepit idea that holds no water. Yet, scientists think that it may have a genetic component. However, we simply cannot find it. I find this very odd and was wondering if there were any studies (as of December 2021) that could shed some light on why some people are gay, straight, bisexual etc.
https://www.pbs.org/newshour/amp/science/there-is-no-gay-gene-there-is-no-straight-gene-sexuality-is-just-complex-study-confirms

Rather than sexuality which definitely has a genetic basis - without which the species would die out, are you perhaps referring to sexual orientation? Could you point to the studies that you claim "disprove" a genetic or epigenetic connection to show us evidence of your prior researches as per our requirements laid-out in the  [help]. Please also take our [tour].

Certainly. https://journals.plos.org/plosone/article?id=10.1371/journal.pone.0014821

Comments are intended for discussions that lead to improvements in a post. They are ephemeral and often overlooked; consequently they should not be used to convey information important to the post. Instead please [edit] your response into your post. ——— 2) Please give full reference information not just a link as links break. This is a good example of how to format references. ——— 3) Note that the paper you linked isn't about sexual orientation and their failure to find epigenetic markers is not proof that such markers don't exist!

It was cited as such. I didn’t read the full article.
I think you are being a little draconian. This is my first time posting on stackexchange biology

For what it's worth, there is never such a thing as a "something gene", that's just not how it works. All humans have the same complement of genes and this is a classic misconception in media reports on genetic issues. Can you explain what it is you say that "scientists" concluded in 2019? Can you also explain how the paper you linked to is relevant? That seems to be a study on ageing, and specifically how epigenetic modifications change with age, so I don't see the connection to your question.

Added source. The source uses the term gay gene.

As currently written your post suggests you lack the background to understand a real answer. Your source says "There is no gay gene" — what do you think you are supporting by adding that source? Science is a process not a body of "facts" — you will have a much better experience here if you do some additional reading and make sure you understand the basics of genetics (or if you do have that background, then make that clear by not using pseudoscientific terms like "gay gene"). In addition, you really must provide references for strong claims such as your second sentence.

there is no one gene controlling X, and X has no genetic basis are two very different things. height has no single gene but it certainly has a genetic component. there is also such a thing as triggered genetic elements, a male has all the genes to make a female and a male, but only one set of genes gets triggered. many genes are like this, everyone may well have the genes for every sexual orientation just as they have the genes for testes and ovaries.

Take a look at Greg Cochran's hypothesis.

On the epigenetics of sexual preference, please see the following:

Balthazart, J. (2020). Sexual partner preference in animals and humans. Neuroscience & Biobehavioral Reviews, 115, 34–47. https://doi.org/10.1016/j.neubiorev.2020.03.024

Balter, M. (2015). Homosexuality may be caused by chemical modifications to DNA. Science. https://doi.org/10.1126/science.aad4686

Wikimedia Foundation. (2021, November 2). Epigenetic theories of homosexuality. Wikipedia. Retrieved December 11, 2021, from https://en.wikipedia.org/wiki/Epigenetic_theories_of_homosexuality.

First article is great. Thank you very much!

You are correct. The Nature article to which you refer does contain the line, "There is no gay gene." But that can't be, and is not, the conclusion. In context:

The largest study to date on the genetic basis of sexuality has revealed five spots on the human genome that are linked to same-sex sexual behaviour — but none of the markers are reliable enough to predict someone’s sexuality.
The findings, which are published on 29 August in Science and based on the genomes of nearly 500,000 people, shore up the results of earlier, smaller studies and confirm the suspicions of many scientists: while sexual preferences have a genetic component, no single gene has a large effect on sexual behaviours. “There is no ‘gay gene’,” says lead study author Andrea Ganna, a geneticist at the Broad Institute of MIT and Harvard in Cambridge, Massachusetts. Ganna and his colleagues also used the analysis to estimate that up to 25% of sexual behaviour can be explained by genetics, with the rest influenced by environmental and cultural factors — a figure similar to the findings of smaller studies. “This is a solid study,” says Melinda Mills, a sociologist at the University of Oxford, UK, who studies the genetic basis of reproductive behaviours. [Emphasis mine; punctuation changed for formatting reasons.]

In stating, "There is no gay gene", you commit the offense that many journalists do when quoting scientific study results: you cherry-picked the sensational sentence, which misrepresents the entire paper.
The second paper you use as a source simply summarizes the original paper, which states,

Same-sex sexual behavior is influenced by not one or a few genes but many. Overlap with genetic influences on other traits provides insights into the underlying biology of same-sex sexual behavior, and analysis of different aspects of sexual preference underscore its complexity and call into question the validity of bipolar continuum measures such as the Kinsey scale.

People cherry pick for several reasons, some innocent, some less so. In 1999, Gur et al published a study on white and gray matter distribution in men vs. women doing certain tasks, revealing a difference. The result: headlines shouting that men really were different than women, and that men had more of the matter associated with logical processing than women. This was a complete misrepresentation of the findings, ignoring that this only pertained to brain usage during certain tasks. Some people used this oversimplification to support their bias that men were smarter than women.
Interpreting scientific articles is sometimes very complicated. Any attempt to boil it down to one take away - other than "it's complicated" - is usually wrong.

I read most of the article. Yes it’s more complicated than just saying “there is no gay gene” but when you are done reading; to sum it up; yes, sexuality cannot be determined by one singular genetic component.

@Aaron Of course the important context is that very, very little can be determined by one singular genetic component, beyond the molecular level. The examples that can are quite unusual and highly notable; that's what the authors are expressing: they have not been able to identify one of those notable, unusual exceptions.

@Aaron - It takes more than 100 genes to determine hair color. So, I would be correct in saying, "There is no single gene responsible for hair color." That is very different than saying/proclaiming, "There is no hair color gene!"