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Non-descriptive gene nomenclature
I formally come from a physics/physical chemistry background, but I have begun to specialize in the area of the biomedical sciences and biomedical engineering. I found myself reading an article which talks about using leukocytes as carriers for targeted cancer drug delivery. One of the frequent adhesion molecules that comes up in the paper is E-selectin, whose wikipedia page I looked up to learn more about the molecule.
On that wikipedia page, it says that E-selectin is encoded by the SELE gene. Now, I can see that the gene is called "SELE" as it abbreviates "Selectin E", but my question is: how did this become the "official" name for the gene? Why doesn't the official name reveal anything about the actual nucleotide sequence of the gene? If someone had never heard of E-selectin before, how is he/she supposed to learn anything about the gene just from the name "SELE" ?
Sorry if this is a very nit-picky question. It's just that most of the substances talked about in chemistry have official names that reveal information about the structure, even if there is a more commonly used "nickname."

Biology names, especially proteins have been a free for all. The investigator that discovered it would usually name it, but there wasn't really any standard. Some named them by molecular weight, such as p21 or p53 which were 21 and 53 kD respectively. Even worse is that people studying different organisms would discover a protein and name it and it was only much later on that it was realized that the molecule had homologs in other organisms with different names. That is how you can get the Son of Sevenless or Sonic hedgehog genes or more descriptive names like DNA ligase or Katanin.

Often times we knew the proteins long before we determined the genetic sequence. With alternate splicing, you can end up with different proteins with different functions originating from the same gene. We estimate there are over 100,000 different human proteins, but only 20 to 30 thousand discrete genes. That means that some of this proteins are fusions of multiple gene products are the result of alternately spliced DNA. So the gene may have been named for a different protein and it was only later discovered that a different protein was a different splice form of the same gene.

Welcome to Biology.SE.
To my knowledge there is no good rule on how to name genes. The first person to name a gene or an allele is free to just pick up a that (s)he finds easy to remember and eventually intuitive. Sometime the author just picks a funny name. Here are a few examples from Drosophila melanogaster data base:

bag of marbles: involved in oogenesis, spermatogenesis.
breathless: involved in development of trachea.
buttonhead: regulates segmentation of the head.
dissatisfaction: involved in many aspects of sexual behavior.
hobo: a mobile genetic element.
Indy: for "I'm not dead yet"; this mutation makes the fly live twice as long.
windbeutel: German for "cream puff".

Did you suggest to name genes based on their sequence? Would be quite painful! One cannot name a gene based on its sequence as the sequence is way too long. The median gene length in humans  is 24,000 base pairs (Fuchs G et al. 2014). You just need to refer to a database to match a sequence to a gene name (see BLAST for example).
How could we improve this system?
Do you have any suggestion on how we should name genes? Should they be named in relation to the phenotype they affect or to the biological pathway that are involved in or maybe something else? The issue is that we often annotate and name a gene before knowing much about its effect.
Note that some genes have several names which is indeed confusing. Would be good to solve this a little bit.

Thanks for the detailed answer! I wasn't suggesting that each gene by identified by its sequence because I know the lengths that we're talking about here. I was just hoping that a kind of "language" would be developed, where sequence could be conveyed in a few letters. I can't think of any good ideas off the top of my head, but let's look at a chemistry analogy: take for instance diethyl ketone. It would be cumbersome to specify an origin position and then provide coordinates for each atom. However, those 13 letters are enough to tell you it's (C2H5)2C=O. That's what I was talking about.

To add to this... there is a transposon in Drosophila called Baggins and a gene called Smaug.. Both these are characters from the novel -The Hobbit

@MuhammadKhan Biological macromolecules are hundreds to thousands of amino acids long Genes can be a few hundred to well over a million nucleotides long, and they are interrupted, meaning that there are long stretches, introns, that get spliced out of the mRNA that makes the protein.   You are asking more of an information theory question in How can we compress the sequence into a form that is manageable but can be decompressed into the full sequence, and that would be difficult. Just take a look at the protein FASTA sequence for SELE and you will get an idea of the challenge.

This is an example of a rather simple biomolecule, Acetyl Coenzyme-A, and its IUPAC name. S-[2-[3-[[(2R)-4-[[[(2R,3S,4R,5R)-5-(6-aminopurin-9-yl)-4-hydroxy-3-phosphonooxyoxolan-2-yl]methoxy-hydroxyphosphoryl]oxy-hydroxyphosphoryl]oxy-2-hydroxy-3,3-dimethylbutanoyl]amino]propanoylamino]ethyl] ethanethioate You can start to see the problems that biologists would face if they tried to adopt chemical nomenclature.

@AMR You're right in saying that the question boils down to something more of information theory rather than biology. I guess I'm just surprised that biologists and information theorists (or people at the intersection of these two) haven't made efforts toward this yet. For your chemical example, I know that names get cumbersome which is why names like acetyl co-A were invented, but at least there is a formal name one could analyze if need be. Gene names don't have "standard" names

@MuhammadKhan people have made effort to systematically name genes.. See an example in my answer below. But they do not make things easier.. It is easy for most people to remember a name rather than a random alphanumeric combination.

Unfortunately there are no rigorous naming conventions for a gene. While some are based on actual molecular function or phenotypic effect, others can be totally bizarre names. Some are actually based on the test-tube and purification fraction labels. Some are named on characters from Mythology, novels etc.
I'll add some examples:
SNARE (proteins involved in vesicle fusion): Soluble NSF Activated protein REceptor (where NSF is N-maleimide sensitive factor).
Nanog: Inspired from an Irish folk tale. There is a magical land called Tír-na-nÓg where if one goes, becomes immortal and forever young. So the discoverers of this gene which responsible for stemness of a stem cell, named it as Nanog.
In fact, a lot of Drosophila genes are named after the visible phenotypic effect of the gene mutation. For e.g. Hairy, Wingless, Sex Combs Reduced etc.
Many genes were actually discovered before the genomes got fully sequenced and since these genes were well studied, the idea of renaming them would not have been very appealing. However, yeast genes have been given formal systematic names.
From SGD

YAL001C   first ORF to the left of the centromere on chromosome I (A is the 1st letter of the English alphabet), on the complement or
  Crick strand
YGR116W   116th ORF right of the centromere on chromosome VII (G is the 7th letter of the English alphabet), on the Watson strand

While I agree that absence of any naming convention would not convey any information about the function of the gene, these wild names do make studying biology interesting. Yeast systematic genes names are so uninteresting and hard to remember (at least for me).
However, there are standard nucleotide repositories such as NCBI, ENA and DDBJ which maintain records about different genes. Moreover, there are standard organism specific databases such as FlyBase for Drosophila, TARE for Arabidopsis etc. In these databases you will find the standard names and description of these genes.

Also have a look at this post: Are there more descriptive ways of naming genes and gene interactions?


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Is there a good easy protocol for extracting proteins (enzymes) from fungal mycelium?
I quantify the enzyme (polygalacturanase) activity by DNSA method. To determine endo-polygalacturonase activity I'm searching for a good easy enzyme extracting method. Any suggestions for a suitable buffer?

Production, Purification, and Characterization of Polygalacturonase from Mucor circinelloides ITCC 6025. The authors use a sephacryl S-100 gel filtration column (chromatography), followed by enzyme activity assay and SDS-PAGE.

source: Stack Exchange

license: Creative Commons - Attribution Share-Alike - https://creativecommons.org/licenses/by-sa/3.0/

sort: votes


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Where does fungus store excess food?
Can anyone pin point, as to where and how, the excess food is stored in fungus (mushroom to be specific).
Like plants store as starch granules, so where does mushroom store?

From the Wikipedia article on fungi (under Characteristics -> Shared Features):

They have a characteristic range of soluble carbohydrates and storage compounds, including sugar alcohols (e.g., mannitol), disaccharides, (e.g., trehalose), and polysaccharides (e.g., glycogen, which is also found in animals).

all_licenses: Creative Commons - Attribution Share-Alike - https://creativecommons.org/licenses/by-sa/3.0/


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Why is absorbance at 280 nm for protein solution going up when I measure repeatedly?
I have been measuring my protein solutions' concentrations by diluting them in water 20 fold with a final volume of 100 uL and then measuring the absorbances of these solutions in 96 well plates with plate reader. I don't remember having any problem up until today.
I used 20 mM phosphate buffer instead of water to dilute them today and measured the absorbances at 280 nm repeatedly three times and the absorbance for one of the solutions went up from 0.043 to 0.068 (absorbance of 20 mM phosphate buffer is 0.030 at 280 nM with same volume); I stopped measuring after third one but it would probably go higher as I measured until I hit a plateau.
I measured absorbances of two proteins and only one of them went up that much, other one went up from 0.071 to 0.088; if this were to be concentration dependent I would expect the second solution go even higher but it didn't happen.
I know there may be differences in UV absorbances if protein is folded or unfolded; would it be that dramatic? What is the reason for that increase? I will be grateful for an explanation and a practical solution to the problem.
NOTE: I increased the total volume to 200 uL by simply adding 100 uL of 20 mM phosphate buffer into all wells and the signal increase slowed down a lot; there is still some increase though with 0.001-0.003 increments in each measurement.

Could it be a slow precipitation?

I don't have any precipitation in stock solutions and I think it wouldn't precipitate after I dilute them.

Is the stock solution in phosphate buffer or water? Depending on the protein, just changing buffers can induce folding changes and aggregation.

It is in phosphate buffer in stock solution.

It looks like your protein concentrations are right on the limit of detection of the spectrophotometer, and changing the diluent buffer changed their concentrations. The samples may not have been thoroughly mixed after dilution and before measurement, so the varying measurements may simply be the solution coming to equilibrium. Temperature can also affect absorbance, so you should verify that your samples have equilibrated before drawing any conclusions. If the absorbance of your phosphate buffer is 0.03, I'd try to keep the sample absorbances above 0.075 or higher to avoid getting too close to the limit of detection. Also, make sure your buffer isn't too old or contaminated with something which could be affecting its absorbance characteristics.
I would suggest taking one or two protein samples and doing a dilution series (1:1, 1:5, 1:10, 1:20) in a large-ish volume (say 400 ul each, if you can spare it), vortex briefly to mix well, then measure triplicates of each dilution on your reader, along with appropriate blanks (buffer only). You will see differences between each measurement, but it should be quite small, depending on the accuracy and precision of your instrument.
Measured values will not be exactly the same from measurement to measurement, and it would take a lot more than three repetitions to determine if there was an actual drift trend occurring. Measure your sample plate every 5 minutes for an hour and plot the values (don't just eyeball them) to see if the machine may need to be serviced.

These are really great suggestions but most of the time I don't have much protein. I tried using higher volumes but I don't get much absorbance; they are very close to background absorbance. If I use too much protein I both waste my samples and a huge meniscus forms which prevents me from determining the pathlength. I always pipette them very well to make sure they are well mixed too. But your suggestion for measuring for an hour and plotting a graph is a good idea, maybe UV lamp is too old. Thank you!

Could be protein unfolding or changes in conformation. Absorbance at 280 nm is mainly due to the tryptophan residues, and can change substantially as these residues move from a more hydrophobic (buried inside the protein) to a more hydrophilic (exposed to the solvent) environment. Your protein may be reacting to the new buffer.

According to the OP's comments the protein is being diluted into the same buffer (I think).

The way I understood it, OP swtiched from water to phosphate buffer for this particular experiment, and is getting the result for one particular protein sample in that buffer.

This article explains the effect of protein unfolding on amino acid absorbance with clarity detail: http://www.life.illinois.edu/biochem/455/Lab%20exercises/Photometry/spectrophotometry.pdf

I diluted the protein with the same buffer I it was in. Sorry for the confusion. Thank you for the paper.


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C-myc/anti-myc Ab Interaction with Fusion Proteins
I am going to prepare a c-myc fusion protein with the following configuration:
(28-residue signal sequence)-(c-myc)-(GGSGGGSG Linker)-(Protein of Interest (POI))
POI is a transmembrane protein and the 28-residue signal sequence is needed for membrane incorporation in ER.
My concern is the disruption of the c-myc/anti-myc Ab interaction due to the fact that c-myc being fused on both ends. Does anyone have any experience or reference with a similar construct?

After some further research, I found papers reporting successful co-immunoprecipitation experiments with POI-myc-His fusion proteins.
Moreover, I figured the signal sequence will be cleaved by signal peptidases which will leave the c-myc tag exposed on N-terminus with only an extra residue. I don't think that one extra residue will create a problem.
Here's the tool I used to figure out where the signal sequence will be cleaved:
http://www.cbs.dtu.dk/services/SignalP/
Here are the references for POI-myc-His experiments:

DOI: 10.1016/j.cub.2006.08.085
DOI: 10.1073/pnas.97.2.668


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What is the best way to express two proteins in a mammalian cell?
I have two proteins and I will be preparing a vector with both genes for stable transfection. Each protein will have their own promoter and I will use piggyBac vector to insert a single cassette with both genes in it into the chromosomes. My questions are:

If I use the same promoter (CMV, SV40 or an inducible one) for both genes how would this affect the expression levels? I read in some papers that one of the genes might get expressed lower than the other. If you have first-hand experience I will be grateful if you share it.
If I use the same promoter for both, but put them far away (i.e. ends of the cassette) would the expression levels get affected again? It sounds simple enough but I think people would have published something if they had a good result with that.
If I use different promoters for each gene (e.g. CMV for one and Tet-inducible for the other) how would it work? I couldn't find a paper which used this and I would like to continue with this approach; therefore, if you have any experience I will be really grateful for any help.

I don't want to use IRES or 2A-peptide. I would like to be able to control this experiment in transcription stage.

I was going through my questions and realized this is still unanswered. Well, I have the answer now.
I designed the construct with two separate promoters (CMV for one and Tet-inducible for the other) and got really good results. I haven't had any difficulties in expressing the proteins. I was able to control the Tet-inducible one very nicely and got steady levels of the other protein with CMV.
Long story short: you can safely put two genes under two separate promoters, prepare stable cell lines with them and get good expression levels.

You can use a bidirectional promoter. The problem that you mentioned about proteins not expressed in same level happens because of competition for polymerase. But there are well optimized parts and also commercially available vectors that work fine.
You can clone genes in a serial order. It won't be a problem. Just leave a 100bp linker after the polyA signal of previous gene. If your cassette becomes too huge, then insertion becomes slightly difficult. That's why people use IRES or TA-peptides, etc.
Retroviral based insertions work quite decently, but you can't control copy number variation between different cells.
You can use two different promoters. The dynamics will depend on the promoter strength and concentration of inducer.

Thanks for your reply. I looked into the bidirectional promoters too but decided to try two different promoters. I really didn't want to go with IRES or 2A because I am too afraid to interfere with the interaction, better not risk it :). I will start the experiment shortly and share my experience in here when I get some results.


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Is 1 g/l living biomass for a Biogas fermenter plausible?
A usual number for a healthy biogas fermenter is $10^9 - 10^{10}/mL$ Bacteria, 10-15% of which would be methanogenic archae. Exceptionally healthy fermenters have more total bacteria and up 25% archae.
The mass of E. coli is given with 670 femtograms. If E. coli is representative for fermenting bacteria that would put us on the order of magnitude of 1 g/L living biomass.
This seems like much to me, though this is just a gut feeling.
As additional information, such a fermenter will have a total solids content of about 10%. A typical loading rate will be 4-5 g VS/d l *, two thirds of which get metabolized to gas. To put it another way: The amount of digested biomass per day would be the same order of magnitude as the living biomass. However, as anaerobic digestion is a multistage process, with at least three steps, each population will have a higher rate of digestion to mass.
Are the numbers basically sound? Is E. coli an exceptional bacterium in terms of mass? Are archae generally as massive as bacteria? What are bacteria counts for other fermentation processes?
(*) g volatile solid per day and liter: for each liter fermenter volume, this many g potentially degradable matter are added per day

Not sure if the tag 'digestive system' is really appropriate.

could you write out some of your abbreviations?  a bioengineer might not be here to answer your question, but someone will probably find it for you if someone not from the field could read it.  Also I expect that there is a huge variation in this behavior for archaea.  you want to make a methanogenic fermentor and are looking for an upper limit, yes? this might help...http://www.sciencedirect.com/science/article/pii/S0723202008000222

I think the only possivle abbreviation was VS, did I miss any other?

VS yes; VS/d volatile solids is days- ok. TS, AD?

Fecal matter can contain up to 10^12 microbial cells/gram. The colon is essentially an incredibly efficient biogas fermenter. So your numbers seem plausible to me.

I've done experiments with yeast, and they can easily ferment glucose at the rate of 4g/L/d.  Probably faster, if one were to optimize it.  So your numbers don't seem crazy.

Can you provide a source for that?


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How to produce correct epithet of a species named in the honor of a person?
I want to name a new diatom species in the honor of Prof. Vodyanitskiy. What will be the species epithet: "vodyanitskiya", or other? Are there any rules that regulates how the end of such epithets is changed?

You should check the article 60 of ICN: http://www.iapt-taxon.org/nomen/main.php?page=art60
See in particular Recommendation 60C about specific epithet, and in the specific case, the rule 60C.1 (specific epithet derived from a person name).

You can use the genitive case (as Plant of Vodyanitskiy) as in rule (a), which is independent of generic name, so  if the professor is male: X. vodyanitskiyi, and she is female: X. vodyanitskiyae.
or as adjective (so inflected according the genera), in rule (c): X. vodyanitskiyana, X. vodyanitskiyanus, X. vodyanitskiyanum

Note: there are other rules to have a valid published name, and it is recommended to explain why you choose such name. In additional to the ICN code, there is the companion book: The code decoded. A user's guide to the International Code of Nomenclature for algae, fungi, and plants(Regnum Vegetabile 155), by Nicolas Turland


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E. coli, risk of infection?
There has been a warning about E. coli contaminated water in South Florida. Now I'm wondering are there empirical data or historic cases which show a correlation between E.coli levels in tap water and risk of infection in the population drinking this water. Which pathogens, that could be transmitted through tap water, are common in this area?

This falls within personal medical advice which is off topic as per the FAQ. If you can remove the references to yourself (can a glass washed with warm E coli contaminated water pose a health risk, or better yet, what is the max temperature at which infectious E coli survive), it might be answered, but it's still borderline.

You will know shortly. ;-)

@dd3 Ok as changed it.

I think there has been something of a misunderstanding here between E. coli and the risk of an infection. Normally E.coli is no pathogen, you have a lot of them in your intestines. The measuring of E. coli in tap water and food is a normal procedure to find faecal contamination, since E.coli is easy to grow. But there are other pathogens you have to worry about like Salmonella, Enterococcus, Cholera and also some E.coli. But it is to expensive to test for every possible pathogen. I'm trying to change you question, to what I think you want to know.

@NeoDevlin Your comment above would make the best answer currrently. Unfourtunately, there's no "Not a comment; Should be an answer instead." flag!

The maximum temperature for E. coli to survive is dependent on the strain. E. coli from warm areas can easily survive 45 °C and more. The minimum number of E. coli to ingest in order to become ill is also dependent on the strain. Whether you become ill at all is also dependent on the strain. You harbor many billions of E. coli in your digestive tract without becoming ill, so without further information on the released strain, it's really hard to tell.
One of the suggestions from the WHO Guidelines for drinking water quality is to boil the water in order to desinfect it.


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What organs are absolutely needed by the human body?
The title is my question: what organs are absolutely needed (fatal if injured/removed) by the human body and perhaps animals in general?
I'm not asking which organs are mechanically replaceable, but more what organs cannot be removed (and not replaced) in order to maintain human life.
--update--
To clarify the question, I'm wondering what organs when removed (and applied no medical intervention) would cause fatal effects in a short amount of time, or cause semi-fatal effects (ex. shortened lifespan). A list perhaps of importance, if possible, would be nice.
note: I have no medical knowledge so I'll leave 'short' open to interpretation.

well.. i think it is only the brain which has no mechanical replacement as of now.

@WYSIWYG I've updated my question to clarify. I've been reading in wikipedia that ex. removal of the liver, pancreas is potentially fatal - and was wondering what other organs have the same effect.

I think you can rather ask which organs are dispensable. easier to answer

@WYSIWYG If this is something in dispute or unclear for some organs, I'd prefer to hear what those disputes are surrounding them than change the direction of my question just for easier answering.

Just to clarify, are we looking for organs that once removed would be fatal without medical intervention?  For example the body can be maintained for a long time without kidneys if dialysis is applied.

@RoryM +1 Thanks for the comment. My original intent was in asking what organs when removed would cause fatal effects within a short amount of time(or drastically shorten lifespan), but reading your comment, it would be great if someone could also add in their reply how it would change depending on medical intervention.

As the comments are gradually illustrating, I think this question is too vague to really be answered effectively.  There are some clear examples of organs that can be removed w/o immediately causing death with varying degrees of medical intervention but you could likely keep any small amount of human tissue alive for weeks in the lab so the answer would be all of them except for that piece* (lots of medical intervention required).

@KennyPeanuts +1 Thanks, I see the side question on the effects of medical intervention makes the question itself a bit too vague. I've removed it from the question, hope it makes this clearer to answer.

You can't remove most organs. The heart, lungs and brain are the ones that without you'll die in seconds. The rest you'll live a few hours or more. All depends how long you wish to live. The best way to see which organs are most required is to look at the order organs are shut down when there isn't enough oxygen or blood. That tells you which area most and least important.

+1 Do you perhaps know of a link or source of some sort that covers the shut down list you've mentioned?

Usually when a person has lost blood they'll lose blood to their skin, followed by muscles (excluding heart/diaphragm), gut (from oesophagus to anus). Meanwhile the blood reserve in the spleen is being mobilised, releasing lots of things to help clot the blood etc. Last to go are lymph nodes (involved in the immune system), liver (similarly involved) then heart and brain closely together.

There is a list of "vital organs" on which there is a consensus that if such organs are removed, the body dies within hours/minutes. Yet people can live long with only one lung or one kidney.
Another list could include other organs that are semi-vital: we could live for a relatively long period without them, but their absence has to be compensated with a continuous treatment (pancreas, thyroid, etc.).
In contrast, people can live in "good health" with non-vital organs removed such as legs, eyes, teeth...

+1 Thanks for the reply and vital organs link, is there perhaps also a source you know that covers the semi-vital organs you've mentioned?

I'm not a medical professional, so this isn't exhaustive or guaranteed 100% accurate:
You can live a (*mostly) normal life without your:

appendix
tonsils
spleen
gallbladder
pancreas
portions of your liver (pieces can be removed and it will regenerate)
portions of your small or large intestines
both lungs (one is needed)
both kidneys (one is needed)
uterus
testes
ovaries

*there are some side effects such as lowered immune function or difficulty digesting fatty foods, but for the most part, you might not even notice.
You can live with some **detriment to your quality of life without:

colon
bladder
up to half your brain
stomach
thyroid

** surgical interventions can replace the colon and bladder with pouches of intestine, drugs can regulate the loss of the thyroid. Losing half your brain messes up a lot, but it's survivable.

Can you add at least one reference for this list?

I could be wrong but the question asks which organs are required for life and not with ones are not too important for it.


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Fur genetics - what are the causes of fur colour decision?
This Australian Shepherd puppy's father is a black-tri; mother is a blue merle. What would cause this dog to have so much white color?

Are there any general explanations as to why this might happen in nature? Other examples of this kind would also be greatly appreciated.

One of the reasons can be related to the migration of the neural crest cells that give rise to the pigmented cells. Many genes are involved in this process.


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Does mixing alcohol types result in faster intoxication or worse hangover?
There is a widespread belief that mixing (drink one, then the other) alcoholic beverage types results in faster intoxication and/or worse hangover. E.g. beer, wine, vodka etc. The claim is that sticking to one type is better. Even drinking different brands of e.g. beer is sometimes claimed to cause the same effect.
Is there any truth? Compared to simply computing volume * % alcohol = drunk amount. Assume time to drink that amount is constant e.g. 4 hours.

EDIT:
Thanks guys for the comments and especially @MikeyC's answer! Despite the poorly worded question.
This is a hot topic among youngsters (which I am unfortunately not anymore) but Skeptics.SE (as I flagged my question to possibly be moved to) would have been a poorer fit there as here are all the experts.
Peace.

How can one objectively measure a hangover?  Also, the title is confusing: when you say types of alcohol, I'd expect questions about how for instance methyl & isopropyl compare to ethyl alcohol.

@jamesqf I know nothing of biology. I have flagged the question for moderation attention. But in the meanwhile - edit away, or even recommend me to delete it. By 'alcohol' I mean conventional ethanol.

I think that the belief arise because mixing alcohol types correlated to drinking a lot (and some of bad quality among them) and that correlates to worse hangover. I don't see any chemical base for the belief.

Skeptics.SE requires a "notable claim"; basically, you have to reference something specific making the claim rather than a vague "widespread belief". That said, we don't typically migrate questions that have been answered.

@Vorac: But all varieties of alcoholic drinks have the same type of alcohol: ethanol.   Other common alcohols are toxic, for instance the occasional batch of bad moonshine that containa methanol.  The difference is the other components.  I've seen claims that hangovers are mostly due to the other stuff, not the alcohol.   I don't know if they're more than anecdotal, though, and have insufficient personal experience to judge.

I did a quick literature search for clinical studies, not expecting to find much, but I actually found a published study partially investigating this question using a pretty robust study design. In short, mixing beer and wine didn't seem to impact hangover severity. Since it only covers beer and wine, you'll have to set up your own clinical trial to see how liquor plays with others.
They set out to test whether drinking beer before wine (or vice versa) contributed to greater hangover symptoms, with control groups drinking beer or wine alone. They found no difference in hangover severity between those who mixed beer and wine (regardless of the order in which they were consumed) and those who had only beer or wine. Perceived drunkenness and the occurrence of vomiting were the best predictors of hangover severity. Interestingly, the most objective measure of drunkenness used, peak breath alcohol concentration, was a poor predictor of hangover severity.

Reference:
Köchling J, Geis B, Wirth S, Hensel KO. Grape or grain but never the twain? A randomized controlled multiarm matched-triplet crossover trial of beer and wine. Am J Clin Nutr. 2019 Feb 1;109(2):345-352. doi: 10.1093/ajcn/nqy309. PMID: 30753321; PMCID: PMC6410559.
Full Text Link: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC6410559/

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


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Can plants metabolize iron rust?
It is widely accepted that "throwing rusty nails in the garden doesn't provide nutrients to the plants"(my very similar question on gardening.SE). I was under the impression this is due to iron being in third oxidation state there while biological processes generally use Fe(II). However plants seem to be used to Fe(III)! Perhaps it has something to do with breaking up the oxides?
What is the reason from chemistry point of view for most plants not to be able to break down

hydrous iron(III) oxides (Fe2O3·nH2O) and iron(III) oxide-hydroxide (FeO(OH), Fe(OH)3) (src: wiki)

? Does really none of the paragraph of acids in @Brōtsyorfuzthrāx's answer in the linked question "work"? Assume pH, moisture etc are fine. The soil is iron depleted soil mixed with powdered iron rust.
Let us also assume no steps have been taken to achieve "good" rust.

With limited dissolved oxygen, iron(II)-containing materials are favoured, including FeO and black lodestone or magnetite (Fe3O4).
source

Strategies I and II for dealing with Fe(III) are detailed in the linked paper. Do you have reason to doubt the paper's content? Which part stumped you? Depends a lot on soil pH and other conditions. Can you tell us more what you're after?

Just looked it up, there are over 400,000 different species of land-based plant - you'd get a clearer answer if you've a specific one in mind.

I have changed the word “digest” to “metabolize”, as that is the correct scientific usage. (The uptake and transport stages might not be termed “metabolism “, but they would certainly not be termed “digestion”.)

See this similar question on Gardening & Landscaping SE: https://gardening.stackexchange.com/q/59059/40386

Keep in mind that unless someone has done a controlled experiment supplementing soil with rusty nails and measuring plant growth, any claims about whether or not rusty nails help will rely on some assumptions about soil chemistry and plant physiology that may or may not actually hold in the field. Therefore, a definitive answer might be hard to find.

You appear to have remove the link that answered your question, and now refer and are now asking us to comment on an answer to a question on another list. Whether or not the effect of throwing rusty nails in gardens is widely accepted or not, it is not the concern of a site for biological science professionals and students. It would appear that you have little familiarity with chemistry from your general use of inappropriate terms — originally "digestion", rather than "metabolism", now "break down". I suggest you go back to the expert review, rather than persisting here.

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


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Can Pfx polymerase add only one 3' A overhang?
I am trying to clone a PCR product that was amplified using Pfx polymerase into pGemT vector. I had to A-tail the PCR product using Taq polymerase since Pfx only generates blunt end products. My ligation reaction was successful, however when I got the sequences, there is an extra T between the 3'end-T of pGem and the start codon of my product. It only happens in the 3' end of pGemT (or 5' end of my product). Can Pfx polymerase generates PCR products with just one 3'end overhang? or Can Taq polymerase add two consecutive dATP during the A-tailing?
Please write me your thoughts ASAP, it has happened twice and it's a pain already :(
GGC CGC GGG A-T-T-T-G ATG GGA AGC ATG AAG
The two Ts in Bold belong to pGemT, the T Italic is the extra T I've got from the sequencing reaction. After the T in Italic you may find the 5'end of my insert. The second T (in bold) from pGemT is the one that ligates with the overhang A added through the A-tailing protocol.

Pfx wont add an overhang. Check this. Taq generally adds one A

Could you post sequences at the two junctions, just to make the problem absolutely clear?

Hi, Thanks for your answer! I have included above the sequences at the two junctions as you requested. Hope this helps to better understand the issue.

did you check the electrophretogram of your sequencing reaction. just to make sure that it is not an erroneous base call

Yes, I've checked every peak of the electropherograms and there is clearly an extra T there. From four clones I've sent for sequencing, all of them had this extra T. I have called promega to confirm it is not a vector issue. They said it is possible that they may have produced a wrong batch of pGemT with an extra T. I can't believe that was the case though. Anyways they will possibly send me a new batch next week. If you think that might be a vector issue, please let me know.

Just try with TOPO-TA once.. if you have it in your lab..

Ok, I am wondering if I can amplify the insert from the clone I already have into pGemT? As the coding region of my gene is fine and primers will unlikely pick that extra T, I would like to use the pGemT construct to amplify my gene, because I've ran out of PCR product.

Yeah, as far as the experiment is concerned you dont have any problem. Even with an extra T there is no problem with the current construct wrt expression. You can amplify the clone too if you want. And, it is always better if you keep some sequences ahead of start codon (such as kozak consensus sequences or restriction sites etc).

I understand the extra T won't change the reading frame of the coding region. However, I am planning to subclone this insert into pet30 expression vector. For this step, I need to digest pGemT-insert leaving the extra T in the digested fragment that is to be subcloned. When this happens, the coding region of my protein will not be in frame. Therefore, I will get another protein completely different as the reading frame will be shifted. My question is: if I use pGemT-insert as a template for another PCR, is there any chance to pick that extra T? Forward primer starts from GATG..

I am asking to run another PCR using the wrong construct (with the extra T) as a template because my gene is lowly expressed. I had to struggle a lot in order to get the coding region amplified without mutations. So, running another PCR using cDNA as a template will be a pain. Now that I have the coding region healthy into pGemT with an extra T upstream my gene, can I use it for PCR?


text:
Are these claims of "revolutionising" understanding of human vision and hearing valid?
I've started a hobby machine vision project (and posted some questions to this end on other SE sites) and on a side track, also been looking at relevant research in human vision (and partly, hearing).
I came across the work of one "James T Fulton" through an excerpt of a book which claims to have "revolutionised" our understanding of human vision and hearing.
Here are the respective links:

http://neuronresearch.net/vision/

http://www.neuronresearch.net/hearing/
I tried cross-verifying these claims via Google, but nothing turned up on searching either for the author or the book (apart from some Amazon links and very obscure references).
Considering the controversial natural of the claims made here, I would have imagined some level of debate, but the silence I found on the Internet is puzzling. Puzzling, because the sheer level of DETAIL in these books, prevents me from dismissing the claims altogether as well.
Can someone validate some of the content from these books so I can get an idea of the overall legitimacy of the work (and decide if I should continue reading, or not). Below might be a good link to start from:
http://neuronresearch.net/vision/pdf/11Biophenom.pdf
Adding a few examples:

The simplest one - is that the Principle of Univariance is not entirely correct. This is examined on pages 15 through 17 at this link (where I read it):
http://neuronresearch.net/vision/pdf/11Biophenom.pdf
There is this claim quoted verbatim from the site:
"The theory shows that the ARCHITECTURE OF ALL VISION IS TETRACHROMATIC. Although traditionally called trichromats, it is shown that HUMANS ARE BLOCKED TETRACHROMATS"
Caps in original. I just pasted it here to give an idea of the tone of some of the text.
This specific claim is "proven" here (in short):
http://neuronresearch.net/vision/files/tetracomparison.htm

And a longer form of the text is linked in the same page
There is also a theory proposed on why cochlea are coiled:
http://neuronresearch.net/hearing/pdf/coiledcochlea.pdf

Thanks!

Do you have specific questions? Rather than ask people to evaluate all the claims, it would be better to be specific about what you have questions about. Otherwise, I think this is too broad a question. Generally, if it's not in a peer-reviewed journal, I don't give it a second thought. And even if it is, I still give it a careful scouring.

I do agree. I've posted some examples - but the text is perhaps a bit too detailed. The hyperbole screams BS to me, but I'd rather have some factual debunking of one or two of these theories than just go by the tone of the text. I'm open to further opinion on if the question can be pruned down further or if anything else can be done to make it neater.

Skeptics SE would be a better place for this question because it doesn't directly relate to a biological question. For e.g you said that certain article says that human have trichromatic vision with a suppressed vision. A valid question to ask here would be- how does this work ?
But in your post you are asking us to say whether James Fulton is right or not. It is like asking if HG Wells right about his theories.

I agree. Where I am coming from is - he has stated how it works, and I'm asking if that's a valid explanation (I have updated the question a short while back to this end). If Skeptics is a better place for it, I'd be happy to see it moved. Only thing is I'm not sure the expertise for this question will be available at that forum. Maybe CognitiveSciences? Also, I'm not asking if HG Wells was right - that was science fiction. This is something claiming to be science - I just need to know if it really is. Thanks.

a lot of us frequent Skeptics as well, and as the site is fundamentally about applying reasoning and the scientific method to analyze notable claims, there are a lot of people there that shouldn't have any issue with this. Do you want the question migrated, or would you rather frame it afresh there?

And really if it stay's here, perhaps you could break it down into several questions. For Example: "Why are Tetrapods Cochlea Coiled" tagged with Anatomy, Neurology, and Evolution (maybe more).

Thanks, I think moving the question to Skeptics makes sense if it can be better answered there. Once it is moved, I'll see if the question can be pruned further, or broken down into multiple questions as suggested. Honestly though, even one of these theories getting debunked rationally would be enough, since I'm merely trying to understand if there is any scientific merit  to the arguments this person presents or if most of it is hokum. I just don't have the expertise to judge - hence the range of samples (to pick any one and invalidate - or validate! - it)

To move it to Skeptics, the question should be restricted to a single claim, and preferably that claim should be quoted directly. "Revolutionise the understanding of human vision and hearing" is too vague of a claim for Skeptics. But as the question is answered now, and this would invalidate it, I think it is better to not move the question at all.

I agree the question is high-level but the vagueness was due to the scope of the text and the claims being too broad and the difficulty of mapping the (many, many) claims made to the expertise available here. Pruning might still have been possible if I had further subject matter expertise, to narrow the claims down to exactly where my doubts were. But anyway now that the question is answered, it is moot. Thanks for keeping it here, and it is nice to see the decent secondary responses (votes) etc the question and answer have got. Cheers!

This is a great discussion. I, James T. Fulton, am all in favor of such discusssions However, I note a distinctive lack of citations defending the status quo. The site demands that statements of opinion be avoided; that they be backed up with references or personal experience. I have provided thousands of citations to the relevant laboratory investigations, as well as the relevant mathematical formulations. Please look at the appropriate subset of these applicable to your area of interest and then draw your own conclusions. But please provide citations that you rely upon, not just your gut fee


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Do homing pigeons fly straight home or follow certain "highways"?

Homing pigeons (Columba livia) have been prized for their navigational abilities for thousands of years. They’ve served as messengers during war, as a means of long-distance communication, and as prized athletes in international races. (Source)

Do all homing pigeons just pick a direction, and fly "straight" home, or do they all follow certain "tunnels" or "roads" to get closer to their home area, before heading home? (similar to cars getting on a highway between two regions)
Does any information about that exist? I'd be interested to know if pigeons released in the same area (with different target locations in one common direction) tend to do follow the same paths, or if it's just a chaotic crisscross.

Some time ago I saw a TV programme (probably on BBC in the UK) that claimed they follow literal highways, i.e. they mostly navigate by flying above roads and remembering which intersection to turn off at. (But I've no idea how clearly established this is, given that my reference is a TV programme.)

I would guess visual cues alone would not be enough, as pigeons are rarely transported in an airplane while being able to look out the window. Most of the time they are transported on the ground to the place they have to return home from.

@Nathaniel - Homing pigeons do prefer to navigate along roads, railroads, and other long, straight landmarks, according to Mann et al., 2014 and Lipp et al., 2004

They have mental maps of landmarks, which they use as well as "compass" cues:

... experienced birds can accurately complete their memorized routes by using landmarks alone. Nevertheless, we also find that route following is often consistently offset in the expected compass direction, faithfully reproducing the shape of the track, but in parallel.

--Pigeons combine compass and landmark guidance in familiar route navigation
Landmarks are more important than compass cues in general, and pigeons develop their own landmark maps:

we show that homing pigeons (Columba livia) not only come to rely on highly stereotyped yet surprisingly inefficient routes within the local area but are attracted directly back to their individually preferred routes even when released from novel sites off-route. This precise route loyalty demonstrates a reliance on familiar landmarks throughout the flight, which was unexpected under current models of avian navigation.

--Familiar route loyalty implies visual pilotage in the homing pigeon
That means that different birds do use different routes:

Here, we demonstrate that birds develop highly stereotyped yet individually distinctive routes over the landscape, which remain substantially inefficient.

--Homing pigeons develop local route stereotypy
In fact the same bird might use different routes over time as it learns new landmarks:

a wide intraindividual variability was observed in repeated tosses at the same site; some pigeons remained faithful to the first route, whereas other birds tried successive new routes which, in most cases, were significantly shorter than previous ones. This result indicates that pigeons try, and are actually able, to improve their performance in subsequent releases from the same site.

--Pigeon homing: The influence of topographical features in successive releases at the same site

+1 Great answer. To add to this, homing pigeons are also able to sense magnetic fields (magnetoreception), such as that of earth, to assist them in orientation and navigation: https://en.wikipedia.org/wiki/Magnetoreception

Also solar cues, stars, smell, and (according to at least one group of researchers) gravitational anomalies; and they are able to determine when one of these is being messed with and deprecate the unreliable information.

They use the Earths magnetic field to navigate. <References
Mora C. V., Davison C. V., Wild J. M. & Walker M. M. Nature, 432. 508 - 511 (2004). | Article | PubMed | ISI | ChemPort |>

I study sharks and I have found that several things have electroreceptors like cockroaches, bees, platypus and even pigeons. On top of that. Mammals have what is known as magnetite crystals in the brain. A compass works because it is a weaker magnet than that of the earths magnetic field and aligns with the stronger field. I have a feeling that pigeons use a combination of all of this as well as memory of landmarks and such but I am unsure. I study sharks
https://en.m.wikipedia.org/wiki/Magnetoreception#In_homing_pigeons
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC49775/


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Why does freshly cut grass smell like a watermelon?
I found that, when I smell watermelon aroma outdoors, someone nearby has mowed  the grass. Is there an underlying basis?

I can't say I've ever experienced that, but a cursory Google search indicates this is not unheard of.  Also, a weird Harry Potter fanfic.

Are you saying that when grass is mowed, you smell watermelons, or that when you smell watermelons, someone has cut grass near you?

I meant that the aroma of the freshly cut grass is like a watermelon smell.

cis-3-hexenal is probably the main reason.  It is one of the main chemicals in fresh-cut grass smell (lots of references besides wiki if you google it).  This fairly comprehensive paper on watermelon odorants also says that they find a lot of cis-3-hexenal in fresh cut watermelon.  See tables 3.1, 3.2 and 3.3.

Is this chemical a GLV (Green Leaf Volatiles)

i think so, you can see cis-3-hexenal in this reference on the GLV wiki page: http://www.olfacts.nl/index_bestanden/Page7.htm


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What is an explanation of the dual role of calcium hydroxide in teeth?
Calcium hydroxide induces dentine formation in permanent teeth, while it causes resorption of tooth in deciduous teeth. How is this dual role explained?

For more than 70 years calcium hydroxide has played a major role in endodontic therapy, although many of its functions are now being taken over by the recently introduced material MTA. Calcium hydroxide may be used to preserve the vital pulp if infection and bleeding are controlled; to repair root fractures, perforations, open apices and root resorptions [1].
Calcium ions released from calcium hydroxide stimulates fibronectin synthesis in dental pulp cells. Fibronectin might induce the differentiation of dental pulp cells to mineralized tissue forming cells that are the main cells to form dentine bridges, via contact with cells [2].
Calcium hydroxide ($Ca(OH)_2$) is known to cause internal resorption in the deciduous dentition. Though this has been attributed to chronic inflammation and odontoclasts, the exact mechanism has not been elucidated. $Ca(OH)_2$-induced odontoclastogenesis could be multifactorial. Odontoclasts may result from fusion of cells of monocyte/macrophage series either due to inflammatory mediators or through stimulation by stromal odontoblasts / fibroblasts. Pre-existing progenitor cells of primary tooth pulp because of their inherent propensity may transform into odontoclasts [3].

References:

Carrotte P. Endodontics: Part 9. Calcium hydroxide, root resorption, endo-perio lesions. Br Dent J. 2004 Dec 25;197(12):735-43. doi: 10.1038/sj.bdj.4811897. PubMed PMID: 15608732.

Mizuno M, Banzai Y. Calcium ion release from calcium hydroxide stimulated fibronectin gene expression in dental pulp cells and the differentiation of dental pulp cells to mineralized tissue forming cells by fibronectin. Int Endod J. 2008 Nov;41(11):933-8. doi: 10.1111/j.1365-2591.2008.01420.x. PubMed PMID: 19133082.

Ravi G R, Subramanyam R V. Calcium hydroxide-induced resorption of deciduous teeth: A possible explanation. Dent Hypotheses [serial online] 2012 [cited 2014 Jun 7];3:90-4.

Teeth are made of calcium hydroxyl apatite: $(Ca_5(PO_4)_3 OH)$, which can be transformed by fluoride ions into the much harder calcium fluor apatite $(Ca_5(PO_4)_3 F)$.


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SARS-CoV virus stability at low temperatures
For SARS-CoV-2 and SARS-CoV1 this study published in NEJM by a NIAID team was done at a temperature of 21-23°C (see the Methods section in their supplemental material).
What's known about virus stability in general and for these two viruses specifically at much lower temperatures? (e.g. 4 °C, fridge temperature).

SE Biology is concerned with the mechanisms of biological processes. General information about the coronavirus is available from a variety of reputable sources, some of which are listed here. You are advised to consult them.


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Genetically speaking, are dogs exactly similar to humans and chimps both?
Richard Dawkins mentions in his book The Greatest Show on Earth that dogs are exactly similar to both humans and chimps. Supposing that a cell contains the genetic similarity between 2 species, he says

The human/dog cell should have an identical resemblance score to the chimpanzee/dog cell because humans and chimpanzees have exactly the same degree of relation to dogs. It should be identical, too, to the monkey/dog cell and the lemur/dog cell. This is because humans, chimpanzees, monkeys and lemurs are all connected to the dog via their common ancestor, an early primate (which probably looked a bit like a lemur)

He later says that this will be found within "statistical margins of error".
Because the common ancestor of both chimps and humans was more similar to chimps than to humans, it stands to reason that chimps will be more genetically similar to the common ancestor than humans. Therefore (not a direct therefore), dogs and humans can't be exactly similar as dogs and chimpanzees are, even theoretically. Humans and apes are even more dissimilar. Therefore ape/dog cell will not be exactly same value as the human/dog cell.
Please explain where I am wrong.

Is this premise of your argument correct? - "the common ancestor of both chimps and humans was more similar to chimps than to humans". Since chimps and humans have been evolving for the same amount of time since they diverged, why do you think that chimps are closer to that ancestor than are humans?

I agree a lot with what Alan said.  If I have time I will come back later with a full answer.  At the moment you should read up on genetic distance, which is what we are talking about here.  I doubt if you were to actually calculate it, they would be identical, but you have to go back far enough in mammalian history to a shared ancestor of dogs and humans/chimps. The assertion is that the most recent shared ancestor between all three was the same species.

Physical similarity doesn't guarantee genetic similarity, I understand, but it seems a little intuitive that some genes from a common ancestor pass unchanged in a descendant A that had a similar environment, while changing in another B, that had a different environments, thus making it possible that A will be more genetically similar to its ancestor than B.

maybe you could expand the dawkins quote?  it doesn't really make sense to me.  chimps are much more closely related to us than dogs by most measures.

expanded the Dawkins quote

@AtlLED I accept that it might not be so, but will dogs and chimps have 'exactly' the same degree of relation as dogs and humans? Can we even make such sweeping statements? I think the variation would be noteworthy, and the statement misrepresents the topic.

@Daud even that idea is questionable we don't know what the common ancestor of chimps and humans looks like, some even hypothesize it was an upright animal based chimp knee structure. Also humans ARE apes, human are not dissimilar to apes.

Dawkins is overgeneralizing.  Genetic distance is a fairly new field, and a heated topic, since your answer depends upon which genes you are comparing.  If you picked ONLY strongly-conserved genes for your genetic distance comparison, then you'd find that humans, bonobos, dogs, mice, mosquitos, shrimp, and bakers' yeast are all equally distant (and that the distance is small).  But if you pick ONLY very weakly-conserved genes for your genetic distance comparison, you can get crazy results -- humans are more closely related to yeast than to horses, for example.  (Creationists have had a heyday making fun of some early genetic distance studies that, in retrospect, had poorly-chosen genes.)  So biologists doing genetic distance comparisons try to pick genes that are common to many species, and that have some drift to them, but not so much that the answers are obviously goofy.  Another problem that has arisen is that sometimes an entire gene in several species appears to have been obtained from a very dissimilar species, in a different phylum or kingdom even.  (Current theory is that this is the work of viruses.)  The "tree of life" paradigm might need to be changed to the "web of life."  Needless to say, there's still a lot of argument about which (and how many) genes to pick for genetic distance studies.

Yes and no.  Yes their DNA, like every living thing’s DNA, is made of the same building blocks of A’s, T’s, G’s, and C’s.  But their DNA isn’t exactly alike.   All those A’s, T’s, G’s, and C’s are put together in different orders for dogs and people. You have to understand that 4% with chimps is a difference of actually 35 million single nucleotide differences and ∼90 Mb of insertions and deletions. With a dog only 25% matches up perfectly with a human. I think what he was saying was that all living matter has a common ancestor. From fruit fly to human being according to macro evolution we all came from the same ancestor.
http://genetics.thetech.org/ask-a-geneticist/dog-vs-human-dna
http://genome.cshlp.org/content/15/12/1746.long


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What in neurons and their connections changes during the process of learning?
I'm not sure if this question belongs more in physics or biology (or maybe even computer science)... but biology seemed to fit more.
What changes in the state of our brains when we learn things?  Because I looked on the internet and I learned about artificial neural networks, and every resource I've found teaches of neural networks that have weights that are trained/evolved and then are static at runtime; you simply train the weights and then once you use them in simulations, they never change.
Purely feedforward neural networks that rely on this can't react to new situations differently if they've experienced it before.  They react exactly the same each time.
I suppose that theoretically, a recurrent neural network that was big enough could really learn, but in practice, these have been used as purely memory slots for an existing method defined by the weights, not as storage of new methods of doing things.
So my question is, physically, chemically, biologically, what changes in the neurons and the connections between them when we learn things?  I don't think we really understand how it comes together on a grand scale yet, but I'm pretty sure we've figured out that much.  I want to learn how to model a simplified version of it mathematically/programmatically.

What changes in the process of learning:

The connections (the way one neuron is connected to another). New synapses can form or dissolve in the process of learning. The glial cells such as astrocytes and microglia can facilitate this process.
Strength of connections: Existing connections can be made weaker or stronger in the process of triggering the same circuit. This happens by up/down regulation of the ion channels at the synaptic junctions.

This process is not fully understood and is an active area of research. You can find some mathematical models too for learning processes such as Long Term Potentiation etc.

During learning, LTP (long term potentiation) or LTD (long term depression) can result in post synaptic neurons altering their responses to neurotransmitter release by the pre synaptic neurons. This is achieved at the post synaptic cell by increasing or decreasing the amount of neurotransmitter vesicles, moving them closer or farther away from the axon terminals, and increasing or decreasing the density of sodium channels. All of these factors can lead to changes in the tendency of the post synaptic cell to undergo an action potential as a result of stimulation by the pre synaptic cell, as well as changes in the amount of neurotransmitter released by the post synaptic cell as a result of that action potential. Further, neurons can also establish or degenerate synapses with other neurons, leading to modifications of neural pathways as a result of learning.


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Measuring algae growth or death
My daughter (10th grade) is doing a science fair project on the toxicity of triclosan for the algae Selenastrum capricornutum. She is wondering the best way to measure the effect, given the limited resources at the school.

Should she use the school’s UV/Vis spectrophotometer or just do low-tech cell counting?
If she uses the UV/Vis, does she have to do a calibration curve where she manually counts the cells for some number of data points?
Or can she just compare %absorbance to the controls and before/after numbers?
Or is it already know for this particular algae at a particular wavelength that a certain %absorbance equals a certain number of cells?
If she uses the UV/Vis, does that indicate the total number of cells, dead or alive, or does it only measure live cells?

The trial is over 96 hours.    Thanks!

I've asked some of my colleagues who work on algae to chip in... hopefully one of them will turn up :)

I don't think it's wise to use a spectrophotometer for this experiment, as it is a total count, i.e. it can't distinguish between dead and alive cells very well, and so she might not see a clear difference between her treated and untreated algae. If she has access to a haemocytometer and a good microscope, then she can count the living cells directly, and calculate the cells/ml using a simple formula.
This paper has a nice overview of how to count algae using a haemocytometer, and if her species of algae can swim, then she would need to use the iodine solution mentioned in here:
http://weis.science.oregonstate.edu/files/weis/Protocols/Symbiodinium/Cell%20Counts.pdf
I hope this helps!

I agree with your answer, but it got me thinking. You could use a spectrophotometer if you took identical samples of algae, treated with different concentrations of agent, then inoculated an aliquot of each into a fresh culture (with a fairly large dilution). By reading the OD of these secondary cultures at some later time you could relate the final OD to how many living cells were inoculated. This could easily be standardised by setting up a calibration with different amounts inoculated from an untreated control.


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What is the lipid membrane of SARS-CoV-2 made of?
SARS-CoV-2 is an enveloped virus: in structural diagrams it is drawn with membrane glycoprotein (M), envelope protein (E) and spike protein (S) embedded in a lipid membrane.
What specifically is the lipid membrane made of?
I assume it is derived (somehow) from the host cell membrane, and would have a similar composition.  It doesn't necessarily have the same composition.  If it just host membrane from a typical mammalian cell, it may be mostly phosphatidylcholine (PC) and cholesterol on the outside, and phosphatidylethanolamine (PE), PC and cholesterol on the inside.  It should equilibrate over time, since the virus does not contain a flippase to keep an asymmetrical distribution.
Related questions: Does the composition depend on the cell that the virus was assembled in?  Does it have any selectivity, or does it simply pick a random piece of the host membrane?  How exactly is the viral membrane detached from the host?
Probably most important question: Does the infectivity of the virus depend on the composition of the membrane?

Structural protein (M, also S,E) have transmembrane domains anchored inside cell's Ergic ER and Golgi infection-modified vesicles, they'd take some lipids with them during the assembly of the virions (expression of M plus low amounts of others produces self-assemblying viral like particles), origin of those virion factories vesicles is unclear.

The last reference says it is the virion factory vesicle (1 2 3) which is exocytosed all at once.

I don't know if this questions has been studied in detail even for SARS, let alone SARS-CoV-2. As noted in a comment by reuns, the virus lipid envelope is most likely derived from that of the endoplasmic reticulum.
What we know however from from some studies on influenza viruses, that's hardly the end of the story though:

Using rigorous virus purification, marked differences between virions and host were observed. Over 125 phospholipid species have been quantitated for three strains of influenza (HKx31- H3N2, PR8- H1N1, and VN1203- H5N1) grown in eggs. The glycerophospholipid composition of purified virions differs from that of the host or that of typical mammalian cells. Phosphatidylcholine is the major component in most mammalian cell membranes, while in purified virions phosphatidylethanolamine dominates. Due to its effects on membrane curvature, it is likely that the variations in its content are important to viral processing during infection.

And this part is a bit of a "reverse answer", but lipid rafts on the membrane of the target cells are thought to be important for the entry of coronaviruses, including SARS. One experiment found that depleting cholesterol from the cell membranes dramatically slowed SARS virus' ability to enter them.
(Regarding how SARS-CoV-2 might come out [with different envelopes] from different cells: we only had confirmation very recently that it substantially infects intestinal enterocytes. So expecting a lot of extra details on that is a bit premature right now.)


text:
Are there any enzymes without aromatic amino acids?
I'd like to try a new spectroscopic technique to study enzymatic reactions (which reaction doesn't especially matter, something simple and with fast kinetics like catalase would do fine - I'm just trying to test the technique).
Unfortunately, for this technique to work, it is best if there aren't any aromatic amino acids around since it is entirely in the UV range.
I did a quick search in UniProt, and - somewhat surprisingly - there are very few proteins without aromatic amino acids, let alone any enzymes.  The yeast protein sequences database contains zero(!) proteins with no aromatic AAs.  The human database contains a few proteins (~20): for example many metallothioneins, proline-rich proteins, and HMG proteins have no aromatic AAs.  No common enzymes turn up in that search.
Can anyone think of an enzyme - or perhaps a small catalytically active fragment of one - which has no aromatic amino acids?
If not - this is maybe a more general question - what makes aromatic amino acids apparently necessary for most enzymes?
P.S. The yeast results are:
lowest aromatic AA content is 4% in vacuolar ATPase (many DNA/RNA polymerase subunits are also quite low);
median is 11%, many common enzymes eg alcohol dehydrogenase, glutathione oxidoreductase etc fall in that range;
highest is 24% in sterol methyl oxidase
UPDATE A thorough search of the RefSeq NR protein sequence db does turn up a whole lot of stuff, but it tends to be from weird species, mostly bacteria... eg oxaloacetate decarboxylase from Rubrivivax benzoatilyticus (which - maybe no surprise - eats aromatic compounds).  It still seems like there is a strong selective pressure to incorporate aromatic AAs in enzymes

A control for your assumption of selective pressure would be info on proteins that lack other amino acids or groups thereof. I know that is not your particular concern, but…. And out of interest, is it possible to state what replaces the aromatics in the odd-balls you mention — can you do MSAs? If nobody else has done it you might even be able to publish your results.


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Is it possible to crystallize proteins so that the crystal has a normal concentration of potassium chloride?
I've been learning about protein crystallography, and how crystals are made.  A lot of the crystallization processes involve gradually changing variables (protein concentration, pH, salt concentration, precipitant concentration etc) for example through vapor diffusion in order to drive the process forward.  One of the things I noticed is that it is very rare to see sodium, potassium or chloride ions in the finished crystal structures.  The crystals tend to have a lot of water in them (both in channels between the proteins, and bound to the proteins and visible in the crystal structure) but much fewer than expected visible ions.  Back of the envelope, 0.15M KCl should have one K+ and one Cl- per 11 nm^3 or per 370 waters; in reality there seems to be almost no ions visible in the structures, except strongly bound ions as cofactors like Ca2+, Mg2+, zinc, iron etc. It is hard to tell if that is because the ions are there but not in a periodic structure, or because they are not there; and also hard to tell what the final concentration of ions is.
Is it possible to crystallize proteins so that the crystal has a concentration of potassium chloride similar to that inside of the cell?
Does having KCl (or NaCl) at normal physiological levels prevent proteins from crystallizing at all? (regardless of other conditions)  Or, is this something that is possible but not done usually? (eg in favor of much higher/lower concentrations, or having different ions around)  Or, is this already the usual state for prepared crystals? (but just not visible in the structures because the ions are always disordered)
Additional question: If the effective concentration of KCl in crystals is very different from physiological, does this significantly change the protein structure?  Has there been any study of the change in structure vs type and concentration of ions?


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How fast do lipids on the inside and outside of a lipid bilayer exchange?
Biological membranes normally have different composition of lipids on the inside and outside (ref 1, ref 2).  This is maintained both by how new lipids are added to membranes, and by specialized enzymes (flippases).  Certain lipids being on the "wrong side" is an important signal (eg for apoptosis).
My question is: how fast do lipids from the inside and outside exchange spontaneously?  In a patch of membrane with no enzymes, how long until half the "inside" lipids have flipped to the outside and vice versa through simple diffusion?
I did a search but couldn't find any specific paper that measures the spontaneous exchange rate.  I'd be very interested in a definitive answer - could be just order of magnitude, but based on real measurements.
UPDATE This is important for figuring out the inside/outside composition of membranes in real scenarios when they're no longer being maintained in an asymmetric state: for example enveloped viruses, but also microparticles, dead cells, etc.

A cursory search seems to show at least a few papers using sum-frequency vibrational spectroscopy to measure rates of flip-flopping/transverse diffusion in lipid bilayers.
This paper in particular ("1,2-Diacyl-Phosphatidylcholine Flip-Flop Measured Directly by Sum-Frequency Vibrational Spectroscopy") cites an activation barrier of 79 kJ/mol which corresponds to a rate on the order of 10^-4/s.
There is also this review which seems to have more information on the topic: http://dx.doi.org/10.1021/acs.accounts.6b00435.

Very well. Also, this paper may be helpful, as it has some data about the activity of Mdr2, a phosphatidylcholine flippase by the measurement of time and relative fluorescence . One may also find the rate of movement implicitly .   https://www.sciencedirect.com/science/article/abs/pii/0092867494904464                                       @Spencer Guo by the way, the first link of yours is not working.

@SpencerGuo - Good stuff, thanks.  From the first paper, looks like at human body temperature the half lifes are < 1 min for DMPC, 9 min for DPPC, > 300 min for DSPC.  Very dependent on lipid tail length...  The review is also excellent.

@AlexI Yes I should have noted that the barriers are quite strongly lipid identity-dependent.


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Are there any small peptides with protease activity?
I'm looking for a very small protease to base an engineered protein on.  I did a search for "peptides with protease activity" but all the results seem to be as peptides to be used as protease substrates (eg in assays); I couldn't find any catalytic peptides.
What is the smallest protease or domain which has independent proteolytic activity?  (Ideally looking for something very short, a dozen aminoacids)

60 residues is the smallest I could find, ref below.
https://jbiomedsci.biomedcentral.com/articles/10.1186/1423-0127-19-54

feel free to edit answer for style


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Protein crystal X-ray diffraction at room temperature?
It seems essentially all protein X-ray diffraction structures are obtained with flash-cooled protein crystals (in a stream of very cold gas).  I’m curious if the diffraction pattern would be significantly different from the same crystal at room temperature (from thermal expansion, but not only - it may favor different conformations, and also flash cooling may disorder parts of the crystal).
What are some of the reasons that X-ray diffraction uses cooled crystals?  Is it possible to do the same experiment with a room temperature crystal?  Has this been done before, and what happens?

I don't know but I think it would have something to do with vibrations in the structure at room temp vs low temps. A la Brownian motion.

Please read the help on asking questions. You will see you are required to do some research before posting and show evidence of this in your question. A simple Google search ("why are crystals cooled for protein x-ray crystallography?") gives an immediate answer. And asking whether anyone has tried it at room temperature. Really!

@David Why not answer the question rather than try to police question quality?  I did do a number of searches; I just did it on pubmed, and the results were all about how to make crystals.  I also looked through recent structures on rcsb.org and they were all cooled.

Because it lowers the quality of the site. As an educator I try not to feed people with answers, but teach them how to answer questions themselves, even if they do not thank me for it.

see also Science (2018) Cooling Crystals is Great. Except When It Isn't.

While protein diffraction data is nowadays most often collected in cryogenic temperatures, the room temperature crystallography also exists and you can find thousands of datasets in the PDB that were collected in 300K.
The main reason for cryo-cooling is that it slows down radiation damage.
You can find in-depth discussion of both options in the literature. I just quickly googled it and found two review articles about advantages of cryo- and room temperature crystallography:
Cryo-cooling in macromolecular crystallography: advantages, disadvantages and optimization
Macromolecular room temperature crystallography


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Which algae grow fastest under optimal conditions?
I'm interested in what the fastest growing microalgae are under optimal conditions for photoautotrophic growth (energy from photosynthesis only).  Fastest can be interpreted as either shortest doubling time, or maximum biomass yield per day per unit area.
Many of the algae investigated for biofuels are fast-growing, for example Nannochloropsis and Ankistrodesmus.  There are many publications that give yields in pond culture or in outdoor photobioreactor, for example Nannochloropsis at 20g/m^2/day (dry mass).  However it is not as common to see yields for lab photobioreactor (axenic, possibly with continuous illumination, and all other conditions optimized for fast growth).  I'm mainly interested in what family of algae would be evolved for fast growth, and where they may be found in nature.
EDIT As one of the answers pointed out, algae can grow heterotrophically as well and that may not be a fair comparison.  I changed the question to be about autotrophic growth (from photosynthesis) only.

The answer to your question depends on many variables. To narrow down the list of possible species it might be helpful to list under which conditions you intend to grow this algae and whether you want to give an additional carbon source such as glucose or acetate.
To give you an idea why this is important: the fastest growing alga I have ever worked with is Chlorella sorokiniana with double timings in the range of two hours. However this was in acetate containing medium and it would not have relied a lot on photosynthesis to grow nor would this make a lot of sense in open pond cultivation systems.
In terms of photoautotrophic algae, we have seen the most consistent quick growth (OD) in our system (Algem photobioreactor) for Dunaliella species. But this is based more on experience than rigorously testing a lot of species. In general there also seems to be a correlation between size and growth speed. Some cyanobacteria have a growth rate of around two hours Article

Thanks!  You are quite correct, I should have mentioned that I'm interested in the fastest possible photoautotrophic growth.  There could be extra CO2 but not extra organics in the growth medium (well, maybe vitamins but not organic energy sources).


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Ways to monitor enzyme kinetics with very fast time resolution?
I'm interested in any way to do time-resolved study of enzyme kinetics.  I am studying some physical variables that may affect kinetics, but I want to study how quickly they take effect, and how long the effect lasts.  The time scale of interest is milliseconds or less, so the typical spectrophotometric methods (which usually look at absorption change over fairly long times on the order of minutes) won't work.
Luciferase activity can be monitored by tracking the light emitted, and this has potentially very high time resolution as the time between reaction and light emission is on the order of microseconds.  However luciferase is a bit of an odd model, as it is not part of normal metabolism or signalling in most organisms.
Is there a way to track enzyme kinetics of any other enzymes with a similarly fast time resolution?
EDIT Just discovered one name for what I'm looking for: "transient kinetics".

Is the change you want to observe visible in the UV/Vis range and you're asking for a way to observe this with very high resolution?

@MadScientist: I'm looking for good model systems, not any specific enzyme.  The change of many enzymes is visible in UV/Vis, eg anything coupled to NAD/NADPH, unfortunately if doing a conventional progress curve type measurement the time resolution is more on the order of seconds or tens of seconds, not milliseconds.  So it either has to be a somehow sped up way of doing progress curve, or some other way.

Are you familiar with Stopped Flow? That sounds like what you're looking for.

Some ATPases can work with MANT-ATP or similar fluorescent ATP analogs that change their fluorescence properties upon protein binding as well as hydrolysis of their phosphate group. This has been used frequently (see here or here) to study enzyme kinetics on fast timescales.


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What determines if a small protein / large peptide is immunogenic?
I'm wondering if there is some threshold in size or a specific structural property that determines if a small protein or large peptide would cause an immune reaction.
Context: there are a number of drugs being developed based on antibody mimetics (small protein scaffolds with stable folding).  These tend to have a size around 50-60 aminoacids (6-7 kD).  These tend to be much easier to design than human / humanized antibodies.  However, it is not uncommon for these to cause anaphylaxis, especially if administered repeatedly.  It may be good to somehow design a version of a small antibody mimetic which itself does not cause an immune reaction.
Are peptides around 10-20 aminoacids typically immunogenic?  What about slightly larger ones up to 30 aminoacids?  Does it matter if they have a stable secondary or tertiary structure?

Do you mean "antigen mimetics" instead of "antibody mimetics"?

I mean proteins like affibodies or minobodies  - see https://en.m.wikipedia.org/wiki/Antibody_mimetic

Here is a useful article on this, https://www.prosci-inc.com/resources/antibody-development-guide/what-is-an-immunogen/

Fundamentals and Methods for T- and B-Cell Epitope Prediction

Edited to delete off-topic section:
Most (all?) proteins are ultimately immunogenic. You can't really design your way around immunogenicity, which is a good thing, because otherwise we'd all be dead from viruses.
Obviously not all proteins are equally immunogenic, but basically all protein therapeutics are immunogenic at some rate. One thing that seems positively associated with immunogenicity in that review is quaternary structure, but it's quantitative not qualitative.
Some of the commonly used synthetic antigens used in biochemistry such as HA C-terminus (24 residue) or FLAG (8 residue) are quite small.
More generally, this study finds that even 3-5 residue peptides can be sufficient to raise immune responses. So likely any polypeptide is potentially immunogenic.
Such small peptides are unlikely to have meaningful secondary/tertiary structure. My guess is that that you would have to have a pretty structurally strange protein to avoid any potential immune response. However if you had an insoluble or somehow sequestered protein you might be able to avoid it, but I'm not an expert so I can't say how.

Thank you, fascinating info.  I'm looking in the opposite direction, for ways to have peptides not be immunogenic / less immunogenic - so the exact opposite of adding a carrier protein.  Basically the idea is to have engineered small protein as a drug; if the drug itself causes an immune response that's a problem.  For example Ecallantide is a ~6kDa protein drug based on Kunitz domain scaffold; it causes anaphylaxis in about 4% of people who take it.

yeah, i realized that halfway through writing it and had to go back and rewrite it with the knowledge that that was your motivation. basically, as i said above, it's hard to make proteins not immunogenic.

see also wikipedia on immunogenicity: https://en.wikipedia.org/wiki/Immunogenicity. basically, (nearly) all peptides are immunogenic.

@AlexI deleted irrelevant hapten section and added some more refs.

"Such small peptides are unlikely to have meaningful secondary/tertiary structure" - That should be mentioned in any basic textbook (hapten chapter)!


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How to find out what biological effects a molecule has, without having a specific mechanism or pathway in mind?
I'm interested in finding what biological effects a (small) molecule has in a high-throughput and "low-assumption" way. I'm mainly interested in cell-based assays.
Background: There are easy/fast assays of high level phenotype effects eg cell proliferation or differentiation, but those wouldn't pick up lots of molecules as being active at all, unless they're at toxic levels. There are also a lot of assays which look at a specific pathway, eg using a Western blot with an antibody to a phosphorylated protein (phospho-Akt comes to mind). But those don't help if one doesn't already know the relevant pathways; they're more useful if screening molecules that target a known pathway, or confirming a known/suspected mechanism of action (and also most really aren't high throughput).
I'm interested in assays which can produce information on everything a molecule does when interacting with a cell, if possible in a single experiment.
Some things that come to mind:

Gene expression profiling (eg using microarrays or RNAseq): gives information on the expression of everything; it is comprehensive and reasonably high throughput. The main limitation is that expression is usually a few steps downstream of what the molecule would interact with, unless it is literally interacting with transcription factors; so it would usually require tracing changes back up various signalling pathways
Proteomics (eg 2D-PAGE, shotgun proteomics): gives information on protein abundance, and potentially post-translational modifications. Seems hard; for 2D-PAGE for example typically a couple of thousands of spots can be identified (a fraction of the whole proteome), and the amount of protein in each measured to within ~20% accuracy (semi-quantitative). Possibly also needs a fairly large sample
Metabolomics (eg LCMS): gives information on all metabolites

Are there any other types of assays that would give information across all/many genes/proteins/enzymes/signalling pathways?

Okay, I cleared up the comments here including my own. You're right, they're unfriendly. But, on the other hand, your question is pretty rude, too. You're asking for a shortcut to knowing everything, seemingly without having done any work yet yourself. It does sound like you're looking for something very similar to a company recently in the news as a scam. Why do you think there is some magic secret method you can learn by asking a single question on Stack Exchange while everyone else in biology is working hard on other methods? Do you think they're just wasting their time and money?

You're also dismissing techniques that other people put a ton of effort into developing because they're "too hard". Science isn't easy. Everything easy is already done, hard is all that is left. If it's easy, it's not valuable.

@BryanKrause I'm sorry it came off that way. I am actually very familiar with the opposite problem: given a target protein/pathway, find small molecules/peptides/proteins that interact with it. It seems most biochemical and cell based assays are optimized for that, while finding all things that a specific single small molecule interacts with is a much less common type of screen. Possibly it's just my unfamiliarity with that type of workflow, but the answer is highly non-obvious. I don't expect a secret method, and I'm not dismissing methods which are hard, just noting that

Personally I think this is a poor question, because the answer is immediately obvious to anyone who was worked with current technologies for both proteins and nucleic acids, but as always, it is hard to prove a negative.
The short answer is NO, not even close to a single experiment or assay that can tell us EVERYTHING  a single protein can do/interact with. If there were, we would be in a situation where almost all of cell biology could be done in a few thousand experiments.
Let's take a well known protein that people have been studying for quite some time now - the "Guardian of the Genome", Tumour Protein p53 (TP53, AKA p53). This is admittedly an integral protein to cellular function, so has a lot of interactions - but it serves to illustrate the point.
Now, let's head to the STRING database for this protein. STRING is a database that charts protein-protein interactions through a number of sources, mostly from the literature, but also through predicted interactions, with varying levels of confidence. Check out the legend and settings tabs for the page to see how those work.
The initial page for p53 should show you p53 and 10 other interacting proteins - these are the most highly significant interactions, with the greatest confidence in them being real. Now, click on the "+More" tab, and you should see 20 interactions - do this a few more times and watch that number and the complexity of interactions grow - it'll show interactions, not just with p53 but between other displayed proteins too. This is a fairly simplistic way to show the number of interactions a single protein has inside the cell. You should finally come up with a few hundred different proteins that have been found to interact with p53. Now, don't forget these are protein-protein interactions only.
p53 is also a DNA binding protein - it is predicted to interact with the genome at about 55,000 binding sites (see sect 2.1, para 2 in this publication) - What does it do at each of those sites? Short answer is - we don't know. Maybe upregulates something, maybe downregulates, maybe nothing, maybe something else.
There's also some evidence that p53 interacts with various types of RNA molecules, some of which are only relatively recently discovered, so assays for them are still fairly primitive. Again, what it does there, we don't know.
If you have ever worked with proteins and DNA, you will know that the methods of working with each are usually quite distinct - which also puts a technological barrier to your idea of a single unified assay. Of course, it is entirely possible to do DNA or RNA-protein interaction studies (e.g. ChIP), but to do these at the same time as protein-protein in a single assay - no chance.
There's also a second (third?) problem - sensitivity. Currently, with most technologies, you can have it one of two ways, but generally not both! You can either detect the rare interactions - the 1 in 1,000,000, by removing the common ones; or you can see the common ones but not the rare, because the common ones swamp the rare. An example of this is in RNA sequencing. In a cell, ribosomal RNAs (rRNA) make up about 80% of cellular RNA, despite not coding for proteins (that I know of). So, if you want to look at a cellular coding RNA by next-gen sequencing, most of the time you will do a rRNA depletion or else 80% of the raw signals will be rRNA, which is a waste of your useful information achievable.
Edited to add:
As has been clarified in comments, OP is not asking for a single assay to do all, but rather types of assays that might be suitable to produce similar results - basically things like Next Gen sequencing, but for small-molecule (not necessarily protein-protein) interactions, which essentially turns this into a shopping question, which are generally not considered appropriate for this site as tools/techniques change quite frequently. It might be more appropriate on bioinformatics.
As always, the content I wrote above applies, particularly the bits about sensitivity and technological barriers.
As far as I know, no such assays exist for proteins, but the one that might come the closest is protein mass spectrometry by MALDI-ToF with de-novo peptide sequencing. These are (in my very limited experience with them) by no means as trivial as setting up a Next Gen sequencing run, but I am sure the technology and associated bioinformatics tools will improve and mature over the next few years.

Thanks for the answer; I'm sorry you think it's a poor question, let me know how it can be improved.  I'm looking mainly for a way to find what external treatments interact with; small molecules which act on surface receptors or diffuse inside the cell, rather than interactions of proteins already in the cell. STING is a cool database! I looked at the type of experiments used to build it, a lot of it is either co-immunoprecipitation or affinity chromatography. These are specific to PPI, again very cool but don't work for the small molecule scenario

"a single unified assay" - I mean a single assay per type of result, not single unified assay for everything, of course. For example all proteins a small molecule binds to, or all changes in phosphorylation due to treatment with a small molecule. Roughly in the way that RNA-Seq gives information on all RNA, whereas RT-qPCR only looks at one or a few selected ones.

@AlexI so I'm interested in assays which can produce information on everything a molecule does when interacting with a cell, if possible in a single experiment. doesn't mean a single assay - 'cause that's sure what it reads like to me. You've improved the question, but the complexity of the problem still stands. PPI like with p53 is a single example. Yes, there are mass methods - next gen sequencing is great for genomic and RNA effects, but nothing even remotely similar exists in the protein world that I am aware of.

@AlexI - I also read your question as one assay to determine everything, etc. which, I confess, was the reason for my downvote. That is in reality a preposterous request. If you actually meant something different, it didn't read that way.

@anongoodnurse Yeah, what I meant was one molecule x a view across all targets/pathways/etc, rather than the more usual screen of thousands of molecules x one target. The examples hopefully clarify that, RNA-Seq is an example that works to quantify the effects on the transcription of everything at the same time. I was looking for methods which provide similarly comprehensive data on (eg) binding, phosphorylation, etc. Please feel free to add an answer if you can think of anything.


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Virus-to-virus gene transfer via "sequences integrated into a common host organism"
From "Risks from GMOs due to Horizontal Gene Transfer", by Paul Keese:

In addition to direct HGT between organisms as depicted in Figure 1, forms of indirect HGT have been observed, which involve an additional intermediary organism in gene transfer from a host organism to the final recipient organism. The most notable are virus mediated gene transfer (transduction) between bacteria (Weinbauer, 2004), retrotransfer of a plasmid to a second bacterium, acquiring host genes and returning to the original bacterium (Ronchel et al., 2000), the spread of donor genetic material between several different bacteria coexisting in complex communities or biofilms (Molin and Tolker-Nielsen, 2003; van Elsas et al., 2003; Wuertz et al., 2004); or from virus to virus via sequences integrated into a common host organism.

Does it mean that a viral gene or other DNA sequence gets integrated into the host's genome and then some other virus (that chanced to be in the same organism) manages to obtain this sequence and package it into its own DNA?
Keese's article has no reference for this statement, and perfunctory googling brought nothing to clear the matter for me.

Does it mean that a viral gene or other DNA sequence gets integrated
  into the host's genome and then some other virus (that chanced to be
  in the same organism) manages to obtain this sequence and package it
  into its own DNA?

Yup, that's what it means (although not all viral genomes are DNA-based).
A lot of genomes contain extensive content that is thought to arrive from viral transfer. Here's a reference to get you started:

Endogenous Viral Elements in Animal Genomes (by Aris Katzourakis , Robert J. Gifford; 2010; PLoS Genetics)

If much of an organism's genome originally derives from viral transfer, then any subsequent viral transduction represents a virus-to-virus transfer of genetic information.


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A trivial question on the meaning of "relatives" in an article on horizontal gene transfer
Am I right in suggesting that by close relatives the author meant closely related species, or did he have individual organisms in mind?

DETECTION OF HORIZONTAL GENE TRANSFER
There are several approaches to identify genetic changes due to HGT,
  including:
(1) experimental evidence, whereby a genetic marker is monitored for
  gene transfer to a recipient organism;
(2) phylogenetic analysisof gene sequences to identify topological
  inconsistencies between different gene families;
(3) nucleotide compositional analysis to identify any gene that has a
  nucleotide pattern that differs significantly from the overall genome;
  and
(4) evolutionary scenarios to explain the patchy appearance of a
  genetic signature, sequence or function that is not shared by close
relatives.

(From "Risks from GMOs due to Horizontal Gene Transfer", by Paul Keese)

You are correct.
From the author's point of view there is not much difference, however. Phylogenetically speaking 'species' are kind of artificial distinctions that exist only as sets of individuals and don't usually convey information, so it makes sense to forget that species exist when talking about HGT. Hopefully that helps provide context for why such imprecise wording is so common.