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Does homeopathic or herbal treatment of cancer have any scientific recognition?
Even though we have a very high tech society, cancer is still a serious issue. We humans still are not entirely capable of fighting cancer.
Radiation and chemotherapy are still considered the best methods for treating it and frequently, even these don't work. And, as for this disease, it is a phenomenon of uncontrolled cell growth that has lead to the death of many people. Within these main two treatments, there is plenty of controversy surrounding applying radiation and chemo in a proper sequence.
However, there are also some different methods known and available on the market to treat cancer. Among them, homeopathic and herbal treatments are the most common. So, my question is, do they really work and is there any scientific evidence of that?  If yes then what is it?

Actually modern treatments do not fail most of the time. They most often fail in metastasized tumors - if you catch the primary tumor early enough it is usually removed and the patient can be completely healed. Additionally there is medicine (everything which has an effect) and there is pseudomedicine (everything which claims to have an effect but cannot proof it. Homeopathy and herbal medicine (unless you isolate some pharmacologically active agent from the plants) belong to the last category and don't have any scientific foundation.

While the question is not off-topic, you'd probably make a better fit (and get better answers) on skeptics.SE.

The methodology behind homeopathy is scientific nonsense.  If you dilute anything a billion times, it will have no chemical effect, not even if you shake it all the while.  So no, homeopathy does nothing for cancer, or any medical condition at all.
Of course plants can have active compounds in them, once scientists have identified those compounds, they can be tweaked to make them act better in the human body, and then manufactured under strict guidelines to ensure purity, and tested to determine the optimum dosage; then it's just medicine.  Taxol comes from yew trees, it doesn't mean that native herbalists were successfully treating cancer with it for hundreds of years.  Native herbalists were using willow bark, which contains salicylic acid, which has anti-inflammatory properties, but chemists improved on it, reducing some of its negative side effects, by modifying it to acetylsalicylic acid.
Millions of people get cancer.  Doctors get cancer.  Herbalists get cancer.  If curing cancer was as easy as mixing some leaves in a poultice, people would do that.

The worst thing is that a lot of these quack doctors explicitly tell people that the conventional (western) medicine is dangerous and only their medicine works. While a lot of western medicine is pushed to market by profit-driven drug companies, still a significant proportion are actually effective. In contrast, no homeopathic treatment can ever be effective, and herbal treatments are in general less effective than maintaining a healthy lifestyle!

The short answer: no.
The for-profit pharma companies that so many people love to slam for profiteering? On the other side are insurance companies who want to pay the absolute minimum they can. If there was a cheap plant or mixture of plants that scientifically showed significant success in stopping cancer, the insurance companies would demand its widespread use and stop paying for pharmaceuticals.
Homeopathy is (with few exceptions) ineffective at best and downright dangerous at worst. Most "homeopathic medications" are unregulated (at least in the USA) and unscreened - you have no idea what's going in them, and you have no guarantee that the content is what the bottle says.
A few years ago there was a study done on herbal alternative medicine and the findings were that plenty of these supplements didn't even contain what was advertised. The worst offenders actually substituted known toxic plants for the claimed product.
Even if the product is genuinely what's advertised, there can be processing and purity problems. Recently there was a news article about some people who were severely sickened after drinking improperly processed aconite (wolfsbane) herbal tea bought in a traditional Chinese herbal medicine shop.
The final (rhetorical) question to ask the quacks pushing their homeopathic crap: If ancient medicinal practices are so good, then why was life expectancy back then (and even now the areas where such practices remain in wide use) so low? And if Western medicine's so toxic with all of its side effects, why does the countries that practice Western medicine standards almost always have the best life expectancies?
You can't argue with results - and for all its shortcomings, Western medicine delivers far more than any other known medical practice on the planet.


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Are tongues red because they contain blood?
Many animals have red tongues, though not all of them - blue-tongued skinks and giraffes come to mind. Are they red because of the blood (and therefore haemoglobin) in them?
It sounds plausible, but it also sounds like a post-hoc explanation.
I'd be especially curious about any other species having blue blood, like horseshoe crabs, but still having red tongues, unlike horseshoe crabs, which apparently don't even have tongues.

i would think so, but if a tongue is not red, its probably because there are other colored molecules in the tongue besides iron globins - it wouldn't necessarily mean the blue tongues have less red coloring.

The tongue is just a huge muscle, so it should have its colour from the same source as other muscles - which to my knowledge is blood. Unless it is pigmentated in its skin :) Human tongue I'm pretty sure is due to blood.

Mammalian tongues are red because of haemoglobin. Blue/black colouration is due to the additional presence of melanin.

At the boundary between the epidermis and dermis are melanophores, cells that contain melanin. This brown pigment absorbs ultraviolet radiation from the sun that might otherwise damage the underlying dermal tissue. Melanin in the dark tongues of giraffes (Giraffa camelopardalis) protects the tissue from sunburn as the animals lift their heads into the top branches of trees to forage for leaves. In humans, synthesis of melanin increases upon exposure to sunlight and produces tanning. Extensions of melanophores reach into the epidermis where they inject their pigments into developing hair cells. In a few cases, such as the bright ischial callosities of baboons, pigmented patches of skin are used as visual signals. The blue and red coloration on the scrotum and perineal region of male vervets (Chlorocebus aethiops) is used in dominance displays.

From: Mammalogy: Adaptation, Diversity, Ecology by George A. Feldhamer, Lee C. Drickamer, Stephen H. Vessey and Joseph F. Merritt (2007) p 97 The Johns Hopkins University Press; 3rd edition edition

source: Stack Exchange

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

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What is this plant which looks like "marjoram"?
Place is India.
We are confused about the identity of this indoor plant. My friend is saying plectranthus, possibly barbatus.
I am saying it is marjoram.
Which is correct name for this plant?

It resembles very closely to Marjoram.

This does not look like Marjoram! Marjoram does not have denticulate leaves! Plectranthus sp. has, though. It does not  appear to be P. barbatus either since it has some kind of structures on the leaves that are not visible in your image. Could be another species of the genus ... Please do not consume unless you are sure what it is.

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


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what species moth is this?

found in garden in wiltshire, england. about 15mm-20mm long. Resting at 6pm on raspberry plant.

You have found Smerinthus ocellata, the Eyed hawk moth. It is name comes from the eyes on the hindwing, which do not show at your photo. It is known to show these eyes if disturbed, to scare of attackers.
Photo from wikipedia commons:

The forewing shows considerable variability, as you can see here

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


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What are these Bahrain insects living in a colony?

These insects are in the desert in Bahrain. About 500 in one place both adults and juveniles. Some dung beetle near by. Does anyone know a name?
The adults seem to be protecting the young ?

Welcome to Biology SE! Kindly explain your question further - is it about species identification or what? Currently its hard to get you I am afraid!

Those are hemipterans (true bugs). It is not uncommon for hemipterans to cluster together.

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


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What type of bug/insect is this?

I found this bug on the bedsheet and can’t figure out what it is. Can anyone help?

Could you please edit your question and add the size of that insect as well as a geographical location and season?

It looks very much like a webspinner, Order Embioptera. But many geographic locations would rule that out. I have never heard of these entering homes. I can assure you that it is not a bedbug.

Using my browser's zoom function to magnify the photo we have lets me confirm the order as given (and as the animal has wings, we can be pretty sure it's a male); going to family might be possible for Gina if she still has the specimen, but identification further than that may not be possible -- turning a search engine loose on the order name and "Ross" will (I believe) eventually turn up a notice to the effect that the specialist just named has over 1000 species not yet formally named in his collection ...

It looks like a species of earwig. I get them inside my house at certain times of year or after it rains. As far as I know all species are harmless but I would double check that.

Could you please explain, why you think this was an earwig? For example, the wings of earwigs in my area have quite degenerated wings, not such prominent ones as in the given picture. But this might be different for other species. Would be nice, if you could clarify this.


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Is it Bufo bufo?
I find a couple of frogs. It was cca. 4inches long and 2inches wide. Photo was taken in Prague, Czech republic, middle Europe.. Same zoomed.

Just want to be sure, guys.

I suggest this is definatly a toad. Thank you guys anyway.

Yes, this is almost certainly Bufo bufo.
It has the stocky figure and the warty skin, ruling out the true frogs.
It lacks the stripes of B. calamita or Pelobatus fuscus.
It is brown without the green colors of Alytes obstetricans.
A red yellow or red belly is not visible at the photos, but I assume you would have noticed it if it was present (ruling out Bombina sp.).
See here for some pictures of European toads (not complete and in Dutch, but pictures and latin names are clear.
Finally: a city like Prague lacks specific habitats of the more rare species.  Common toad is the most likely to be found there.


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What kind of a fly is this?
I have a couple of new pictures of insects which I am really curious about. This is one of them. I found this fly near my cottage in Prague, Czech Republic.

It is fluffy with very long proboscis (almost half of an inch in length).

That is in the family Bombyliidae. https://en.wikipedia.org/wiki/Bombyliidae

This is indeed a fly from the genus Bombylius. However, I think this is not the common B. major, but Bombylius discolor.
The reason is that your specimen clearly has small dots on its wings. These dots are characteristic of Bombylius discolor, B. major lacks these dots. Furthermore, B. major has a black part at the front of its wings, that is very distinct. In discolor, this black part is less pronounced or even hardly visible.
Picture from the Dutch Wikipedia:

https://nl.m.wikipedia.org/wiki/Gevlekte_wolzwever

Thank you for your answer. I am so happy that i finally get the right ID of this nice creature and this still amaze me how much i have to learn in ID of bugs.  :)

I found out that this is on of the most common specious from this family in whole Euroupe called Bombylius major. From wikipedia article (same with picture):

The adult is 14 to 18 millimetres (0.55 to 0.71 in) in length, squat and very hairy, with a wingspan of around 24 mm (0.94 in). It has dark patches on the anterior half of the wings and long hairy legs that dangle while in flight. The very long proboscis is used to feed on the nectar of many species of flower, especially primroses. While its wings continue to beat its front legs grip the flower and its long rigid beak is inserted to collect the nectar.[2] Despite its fearsome appearance, the beak is quite harmless.[2]
Bearing a mimetic resemblance to bees their body is stout and furry, with the top of the thorax being black and shiny and the pile either brown, yellow, or white. They have long spindly legs as well as a long rigid proboscis found in the front of the head. Their boldly patterned wings have a distinct dividing border through the horizontal middle between the dark and clear portions. Their antennae are typically very short and pointed. In the field they will be seen hovering and darting above bare ground or flowers, in an up-and-down movement, accompanied by a high-pitched buzz.


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Book for artists' renderings of extinct species?
I'm looking for a collection, in print or online, of pictures of ancient extinct species (i.e., imagined from fossil remains). Is there a good single resource for this? I'm interested in animals of all kinds, barring dinosaurs, and invertebrates. I'm not interested in recently extinct species but rather the very distant past.

Any clade you are particularly interested in? Are you interested in plants? In animals? In Vertebrates? In mammals? Are you interested in a particular mass extinction event? Past 100,000 years or older like dinosaurs (230-65 millions years) or even older? You don't necessarily have to narrow down your question but you might be disappointed to get drawings of ferns, of gigantic dragonflies or of dinosaurs if you were expecting mamooths, saber toothed cats and dodo birds.

You're absolutely right, I should have been more specific. I'm interested in animals of all kinds, barring dinosaurs, and invertebrates. I'm not interested in recently extinct species but rather the very distant past.

With a quick search I was surprise that I could only find

Animals of a Bygone Era: An Illustrated Compendium
Extinct Species
Extinct Animals of the British Isles
Extinct Birds

It looks like there is an open niche to someone who like biology of extinct species and illustrations!
Note all will be of interest to you. None of them seem to offer good quality computer based illustration but rather pencil drawings.


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Mushroom Identification
I live in central Ohio and found these mushrooms growing on our 10 acre property. What is it and are they edible?

Do you have any picture showing the size? And especially the underside of the umbrella?

In addition to showing the underside of the umbrella, it would also probably be helpful to be more specific about the property, and where on the property they were found. In the middle of a lawn? On some mulch? By a tree or stump? Clustered together?


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Bee Identification, Rochester NY
bee I found in Rochester NY, any guesses what species?

Some kind of Halictidae. You can tell by the pollen-collecting ventral side of the abdomen.

It is a bee in the genus Megachile, in the family Megachilidae, not Halictidae.
Megachile are leafcutter bees, and have scopal (pollen carrying) hairs on their bellies (ventral side of their abdomen).
Most bees in the family Halictidae have scopae for carrying pollen on their legs, though they may have hairy bellies with substantial pollen carrying capacity.


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Does the DNA stick to surfaces we touch?
In case of robbery, the police visit the robbery scene and collect some samples like foot print or any item suspected to have been touched by the robber. The sample is then taken to laboratory for DNA analysis. Does that mean whatever we hold or touch, our DNA sticks or drops on it?

"Transmittable" might not be the right word, but yes, we deposit DNA on things we use/touch.

By your use of the term "transmittable", your title make it feel like you are talking about parent to offspring transmission of DNA.

@Mallam_Bello, it's inappropriate to say that"DNA" drops on the surface we touch. The cells of the body drop

Yes, we leave DNA on everything we come in contact with. Even simply standing, we shed skin cells and hair (although only the follicle has DNA) etc. DNA may also be transferred from clothes to clothes.
You might be interested to read this article :) http://www.forensicsciencesimplified.org/dna/how.html

Yes, and as DNA technology becomes more sensitive, there is the unfortunate effect where it's very easy to detect the DNA of a person who hasn't personally been in the location, or was a long time ago. The amplification of DNA is incredibly efficient now, theoretically needing a single cell

This is not only a theoretical thing - it is possible to amplify a single cell.


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DNA of identical twins
I read an article which says that identical twins have 100% similarity between their DNA, but they have different fingerprints. Does that mean the DNA is different in the fingers? What body features can be used to exactly identify the identical twins?

DNA is not the only thing in biology that determines a phenotype (observable trait, e.g. fingerprint). Far from it.
Traits can be affected by the environment (a very good example, temperature-dependent sex determination), stochastic (random) phenomena - these are know to occur in transcription ("reading DNA") or the decay of mRNA (messenger molecules which instruct protein building) - and by epigenetic phenomena. The degree to which DNA is actively read, or silenced, does not wholly depend on the DNA. In fact, this depends on the exact state of the cell or nucleus. Roughly speaking, cells in your brain (e.g. neurons) and the cells that line your lungs have identical DNA, but they are shaped, look, behave and react differently to the outside world. This is due to epigenetics, think of it as the life history of the cell.
Here's an analogy: reading the exactly same book can have different effects on people. The book is the DNA, its information unchanging, but the person and their exact status are the complicated world of the environment and the cell that is using its DNA for some purposes. PS. Terrible analogy but it may help.
By no means is my list exhaustive. Hope you can see how that identical twins with identical DNA can have a different shade of skin color (tanning!), different fingerprints (subtle and complex differences during digit development), or a different sense of fashion (psychology, cognition, what have you).

You hit on the most important points. However, it is also worth pointing out that identical twins don't actually have totally identical DNA. They split from the same embryo and there they have identical DNA, but during development mutations occur which makes identical twins not truly 100% identical. Very close to 100%, but not exactly.

Very good point. I've had the discussion with peers whether or not to include this point in discussions with a layman on these kinds of questions. Bit of a thought experiment: why aren't homozygotic twins perfectly identical? My gut response is not to bring DNA mutations into the answer. I'm sure you'll agree the contribution of somatic mutations to fingerprints is insignificant. Incidence of cancer in twins is much likelier to be caused by genetic history rather than a lifetime of mutation accumulation. But you're absolutely right, it could influence many (inconspicuous) phenotypes!


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Where does the additional 1 proton cost of oxidative phosphorylation come from?
I've been quite confused by the source of the additional 1H+ cost right now. I know it costs the F1Fo-ATP synthase 3H+  to produce an ATP, and it is also stated that:

ADP3−cytoplasm + ATP4−matrix → ADP3−matrix + ATP4−cytoplasm //
  ADP/ATP exchange is energetically expensive: about 25% of the energy yielded from electron transfer by aerobic respiration, or one hydrogen ion, is consumed to regenerate the membrane potential that is tapped by ADP/ATP translocase.(Wikipedia/ADP/ATP Translocase)

So in some books I find out that they regard the net cost of producing an ATP as 4H+. Here is another statement:

The phosphate carrier, which works in concert with ATP-ADP translocase, mediates the electroneutral exchange of H2PO4-for OH- (or, indistinguishably, the electroneutral symport of H2PO4- and H+). The combined action of these two transporters leads to the exchange of cytosolic ADP and Pi for matrix ATP at the cost of an influx of one H+. (Berg JM, Tymoczko JL, Stryer L. Biochemistry. 5th edition. New York: W H Freeman; 2002. Section 18.5, Many Shuttles Allow Movement Across the Mitochondrial Membranes.)

And my question comes here. Why do we not consider the effect of the process of ADP + Pi → ATP on the membrane potential? It also produces a net cost of 2e-. Putting these together, this will lead to 0, i.e. no additional net cost. Or should I only consider H+ but not e-?
(This might lead to another problem about H2PO4-'s breakup.)

The energy required for the ATP production by ATP synthase is obtained from movement of ions (H+ or in some cases Na+) downstream of their potential (proton motive force). I guess, there is no exact stoichiometric exchange as in case of ion pumps. The rotation of the motor is generated by the movement of protons through a channel. The H+/ATP ratio is estimated using biophysical/biochemical techniques.
Brand and Lehninger (1977) estimated this ratio to be greater than 3.
Turina et al. (2003) recalculated this ratio using constituted liposomes and their estimate is  3.9 ± 0.2.
This is an excerpt from a more recent study on chloroplast ATP synthase by Hahn et al (2018).

The symmetry mismatch between the 14-fold Fo rotor and the
  near-threefold F1 head means that the number of c subunits rotating
  past subunit a to generate one ATP is not an integer. cF1Fo requires,
  on average, 4.67 c subunits, or protons, to produce one ATP. The
  nearest integral numbers of c subunits per step would be 4, 5, and 5,
  equivalent to rotation angles of 103°, 129°, and 129°. Surprisingly
  however, the three conformations are separated by rotations of 103°,
  112°, and 145°, or 4, 4.4, and 5.6 c subunits (fig. S5C). This means
  that the position of the c subunits relative to subunit a in the three
  conformations differs.

The structure and efficiency of chloroplast ATP synthase may be a little different but what I want to emphasize is that there is no exact exchange of protons and ATP.

Thanks! Yes, that the calculations are just estimated from experiments is a fair point. But actually it was some statements from pratice problems when I studied biochemistry that cause some ambiguity, for example the contradictory statements 'ATP/ADP tanslocase adds 1 net cost to ATP production' and 'Pi/H+ transfer adds one'. But I will bear in mind that they're all rough calculations. I really thought the F1Fo-ATP synthase was symmetric (regardless of the change in conformation) just like the honeycomb before I read your answer.


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What do the acronyms in C.elegans neuron names stand for?
In this site, I see a variety of acronymic names for C.elegans neurons but what do these names mean (for example AVAL, AVAR)?

Listing all the names would not be possible and hence your question tends to be broad. However, have a look at this site.

Unfortunately, there doesn't seem to be a very systematic nomenclature. Some of the letters are acronyms while others describe a class of neurons. The letters V and D usually describe the ventral and dorsal counterpart of a pair of neurons. The same goes for L and R (left and right).
This list names all C. elegans neurons and you can see that, for example, the "ring interneurons" or motor neurons start with RI or RM. "Inner labial" or "outer labial" neurons are called IL or OL. Ventral cord motor neurons start with a V and their different classes are called VA, VB, VC and VD plus a number. The classes are determined by lineage origins or historic reasons ;) And there are a lot of exceptions to these rules, because their historic names remain even though we continue to learn more about these neurons.
The example AVAL/AVAR are ventral cord interneurons. They form a class with other ventral cord interneurons called AVB, AVD, AVE. So, the first two letters refer to the main class, the third letter to the subclass. The L or R refer to the left and right member of this pair.
Some friends working in C. elegans neuroscience told me that you basically just treat these letter codes as individual names for the neurons without bothering about their origin and meaning too much.

Quote : "Every C. elegans neuron name consists of either two or three uppercase letters indicating class and in some cases a number indicating the neuron number within one class. If the neurons are radially symmetrical, each cell has a three-letter name followed by L (left), R (right), D (dorsal), or V (ventral). "  link


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How does Sanger sequencing resolve two bands with equal size?
In Sanger sequencing, how can we read the sequence if we have 2 bands with the same size?

I can see now how this would be a homework question, although the answer is dependent on the full context which you haven't given us.  Please read the site's policy on homework questions, which requires you to demonstrate having tried to find an answer. Also please note that Sanger was a (very famous) person and his name is spelled with a capital S (which is also used at the begining of sentences in English).

The band size by itself isn't relevant, unless you are using the same primers for PCR as sequencing.

It happens if your sample contains a mix of two DNA molecules, with different bases at a certain position.
In your case, you would have a mixture of ACTGGCTA and ACTGTCTA.
In real life, the "double band" would be half as intense as the other single bands, as the signal is "divided by two".


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Plasma Membrane Proteins and Cytoskeletal Attachment

Regarding membrane protein functions, which of the following
  statements is CORRECT?
a. Membrane proteins are responsible for both cell to cell recognition
  and cell anchoring and are stabilised by linking through to the
  microtubulele cytoskeletal ﬁbres.
b. A protein that is enzymatically active and membrane-bound will
  function significantly differently to a protein that is enzymatically
  active and cytosolic.
c. Membrane proteins are manufactured in the internal cavity (lumen)
  of the rough ER.
d. Damage to the smooth ER could alter the makeup of the molecules
  that are displayed at the cell surface.

Regarding membrane proteins, what is incorrect in the statement “Membrane proteins are responsible for both cell to cell recognition and cell anchoring, and are stabilized by linking through to microtubules cytoskeletal fibres"?
I believe this to be wrong due to it normally being Actin Filaments and Intermediate Filaments that attach to Integrin proteins in the cell surface membrane.

a. Membrane proteins are responsible for both cell to cell recognition and cell anchoring and are stabilised by linking through to the microtubulele cytoskeletal ﬁbres.

Not entirely correct as you yourself pointed out that integrins attach to actin filaments.

b. A protein that is enzymatically active and membrane-bound will function significantly differently to a protein that is enzymatically active and cytosolic.

Possible but not necessary.

c. Membrane proteins are manufactured in the internal cavity (lumen) of the rough ER.

They are secreted in the ER and modified there too but not manufactured there.

d. Damage to the smooth ER could alter the makeup of the molecules that are displayed at the cell surface.

Seems correct because the statement itself does not claim certainty. It says that the makeup of cell surface molecules (which could be lipids or proteins) could change in response to SER damage.


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How to identify an unknown species from its genome sequence
I am currently using ILLUMINA PE DNA sequence data, which I trimmed (Trimmomatic), corrected (Rcorrector) and assembled (SPAdes). I am now interested in using the genetic sequences from my contigs to identify the original organism (I know it is a bacterium).
I have tried using ncbi's BLASTn on one of my contigs with default parameters, on nucleotide collection database. The contig length is about 360 000 bases long. I let the query run for 30 minutes before stopping, nothing came of it. I am assuming 360 kbp is too long.
I was thinking of maybe using a ribosomal subunit sequence (which are used in phylogenetics) to identify the species, or at the very least find the taxonomic family. Is this approach feasible? How can I identify ribosonal subunit sequence from my total sequence?
If not, what would be the best way to approach this question?

Did you use web interface BLASTn or local instalation? What was the library against which you run your query? How long was the contig? 30 minutes seems long.

Crossposted here https://bioinformatics.stackexchange.com/questions/14487/how-to-identify-an-unknow-species-from-a-fasta-containing-genetic-sequences

I’m voting to close this question because it has been crossposted to another (more appropriate) SE site. It also seems to fit this sites criteria for "homework".

@tyersome Definitely Not homework. This is my first time using a StackExchange platform (other than StackOverflow), did not know of the "no cross-posting" rule. Thank you for letting me know.

One of the reasons for the "no-cross posting" rule is that members of one list may spend time and effort answering a question unaware that the question has already been answered on another list.

So how did you solve it? Blasting pieces of your contigs should give good guesses and https://blast.ncbi.nlm.nih.gov/Blast.cgi should work fine with a 360kb contig (restrict to bacteria).

I would suggest using a tool designed for this purpose such as GTDB-tk (slow but informative), mash (very fast but very reference-dependent), or something similar.


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Numbering of C atoms of pyrimidine rings in nucleosides
For a presentation of my work I wanted to show a structural representation for dihydrouridine with numbering of the C atoms of the pyrimidine ring, according to this numbering convention.
Then I found this Paper, where the numbering in 5-methylcytidine is consistent with this convention but that in N1-methylpseudouridine is not.
After some more research I found this paper (fig. 1 A), where uridine is numbered according to the nucleotide numbering convention, but pseudouridne is not.
So why is the numbering of modified cytidine according the the convention (m1C), whereas the numbering of pseudouridine (which is a modification of urdine) is not?

You would probably be best to ask the chemistry SE.

Welcome to SE Biology. I have answered your question, but can I make a suggestion for improving the way you could have asked it. 1. Your question only showed the numbering for uridine, which made it difficult for readers to know what the situation was with pseudouridine. You should have included a comparative picture — the red dihydro atoms in the diagram you provided are a distraction. 2. Most important, you should then have stated clearly what you thought the numbering convention was that was being applied inconsistently. This might have made you consider whether your assumption was correct.

Although this question should probably have been asked on SE Chemistry, as the molecules are of biochemical importance — and the question is based on a misconception — I will answer it here.
Answer
The numbering of the carbon and nitrogen atoms in uridine and pseudouridine is perfectly consistent. The numbering is for the pyrimidine ring, which is the same — uracil — in each case. The different positions of attachment to ribose (N1 or C6)) results in two different structures (and hence names) for the resulting nucleoside, but does not alter the numbering of the components.

[Modified from https://commons.wikimedia.org/wiki/File:Pseudouridine-synthase.jpg‚ Surundu]
Thus N1-methylpseudouridine has the methyl group attached to the N1:

Basis of numbering
The numbering of the atoms of pyrimidine bases derives ultimately from that of the parent, pyrimidine in which the two (indistinguishable) nitrogen atoms are numbered 1 and 3:

The relation of uracil to the parent pyrimidine can be seen from the IUPAC preferred name Pyrimidine-2,4(1H,3H)-dione and a perhaps more easily understood alternative as 2-Oxy-4-oxy pyrimidine i.e. the oxygen substitutions are at positions 2 and 4 (as shown for both uridine and pseudouridine, above).
The preferred IUPAC name for uridine is:
1-[(2R,3R,4S,5R)-3,4-Dihydroxy-5-(hydroxymethyl)oxolan-2-yl]pyrimidine-2,4(1H,3H)-dione
and that for pseudouridine is:
5-[(2S,3R,4S,5R)-3,4-Dihydroxy-5-(hydroxymethyl)oxolan-2-yl]pyrimidine-2,4(1H,3H)-dione
i.e. The formal nomenclature for the two nucleosides only differs in the designation of the position of attachment of the base to the sugar ring (1 or 5 — emboldened above).


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Does Topiramate work by supplanting Pyridoxal phosphate in enzymes?
I have seen it said that the precise mechanism of action of migraine medicine Topiramate is not known. But I certainly see a resemblance between that molecule and PLP (Pyridoxal phosphate, the activated form of vitamin B6).

A significant similarity is that both have an "offshoot chain" (Topiramate's sulfamate taking the place of PLP's phosphate) where PLP's phosphate is what binds it to enzymes in the first place, and both have a more electronegative element in the middle of what would otherwise be a six-carbon ring, apparently important in the typical PLP enzyme reaction.

I have been wondering if Topiramate supplants PLP in the enzyme glutamate decarboxylase, changing the reaction into a deamination. This would deplete glutamate, causing the pill's migraine relief, and the deamination could even cause Topiramate hyperammonemia.
If it does the same or similar to serine hydroxymethyltransferase, which is also PLP dependent, it could cause glycine deficiency, thus collagen deficiency - and this would even explain Topiramate hair loss.
When you look at it this way, it's frankly hard to imagine the mechanism of action is anything different. Have I stumbled upon it? Or has this been suggested already?

After reading the comments I have 3 reminders: 1) Be NICE, 2) Comments aren't for answers or extended discussion, 3) Voting to close shouldn't be used if you simply dislike a post or think it has a simple answer, especially when the OP shows research effort. Downvoting is more appropriate. @MarchHo I think your VTC comment could be converted to an answer

Comments are not for extended discussion; the remaining un-deleted comments of this conversation have been moved to chat.


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Methods for microbial identification in soil
I'm trying to perform a survey of the life in a soil sample. I want to know what species of bacteria, fungi, and other organisms are in it. I've heard that ITS sequencing and 16S rRNA sequencing are both possible ways of doing this, but I don't know enough about these techniques to understand which (if either) I should use, or how you interpret the resulting sequence file delivered by the lab. Will these techniques do what I want? How do I choose one over the other? And how do I interpret the results?

16S and ITS techniques try to identify organisms in your samples by amplifying short, 'barcode' sequences from each organism's DNA, and using those short sequences to try to identify the organism they came from.
16S can be useful for distinguishing between prokaryotes (bacteria), at least to genus level.  It will not always give enough information to determine exact species though.  ITS can be particularly useful for distinguishing between some eukaryotes (plants, fungi, etc), even down to species level.  If you are interested in both bacteria and fungi you would need to carry out both analyses on your samples.
A different approach is to use shotgun metagenomics, where rather than amplify only the 16S or ITS region, you extract and sequence any DNA you can from the sample, and then try to assign each sequence read to a species/genus by alignment with a reference collection (e.g. MEGAN pipeline) or by kmer analysis (e.g. Kraken).
There are some packaged approaches for dealing with 16S. Qiime2 is popular, and will likely involve some amount of work in python to get it working.  There is an R package microbiome, but I don't have experience with this.  Mothur is a long-standing stand-alone package that is resonably easy to use, and MG-RAST is a web-based version that is also reasonably easy.
ITS analysis is less fully developed as pipelines, but I am sure you can find some in the literature by searching for other people's studies of soil microbiomes.
This review of techniques and approaches might be helpful if you are interested to learn more about the whole genome/metagenomics approaches: http://ccb.jhu.edu/people/salzberg/docs/Breitwieser-etal-2017-Metagenomics-review-reprint.pdf

16S is still one of the most poplular methods of idenifying organisms. The technique is very well stablish and it works well. However, it has some limitations, for example there is a significant variance of the copy number of these gene in the species. Which makes quantification not very reliable.
To decide which tools are better to analyise your results, you can have a look a the CAMI challenge paper. Several tools for different parts of the analysis are compared across labs. The most popular tools are not always the best for your analysis.


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Why are restriction sites palindromic in nature?
Any restriction site given in any fragment of DNA or vector molecule is a palindrome. eg restriction site for the endonuclease EcoRI is
5'...GAATTC...3'
3'...CTTAAG...5'

Clearly, this is a palindromic sequence. Why is that?

I was going to answer this question, covering what I regard as a key point that other answers ignored. However as this is a duplicate I have answered it on the original. You may be interested to look at it there.

Not all restriction sites are palindromes : https://en.wikipedia.org/wiki/List_of_restriction_enzyme_cutting_sites:_E%E2%80%93F#Whole_list_navigation
EcoRI: GAATTC (note that this is not 'palindromic' in the usual sense (e.g.'anna', GAATTC read backwards actually gives CTTAAG), but only if given the G-C A-T binding)
The double strand of DNA is composed of one strand in direction 5'-3' and another into the other direction (3'-5'). Restriction enzymes tend to be homodimers, thus would bind to both sides of the double strand, expecting the same recognition sequence. But as one strand runs into the other direction, the strands need to be palindromic to form a double strand.

EcoRI has a palindromic recognition site in the sense of the term used in molecular biology. Perhaps I'm not understanding your point.

@canadianer  my point is that someone asking this question possibly is not well versed in molecular biology, and might thus be only privy to the non-m.-b. sense of the word: 'Palindrome' = 'A Word that looks the same, if the order of  characters is reversed'

You really need an example of the structure of a complex to justify and explain your answer better. What is the point of. binding to both strands?

bukwyrm gives a good answer for "Why" palindrome: to cut both strands. To expand the evidence, apart from clear demonstration of the current utility of restriction enzymes (they cleave recognized nucleotide sequences), the best proof indicating their evolutionary persistence is the relative lack of palindromic sequences in genomes of a wide variety of organisms.1 The idea is that the presence of endonucleases with palindromic recognition has caused organisms with such patterns to lose out to those that lack the patterns over time.

Thank you so much! I had found out lately too!

So that these enzymes can recognize the sequence on the both strands. It's the mechanism bacteria have evovled to degrade the only double stranded RNA and not single stranded.

Welcome to Biology.SE, Sujeet. Your answer could use a little more detail, and ideally a reference to an authoritative source.


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How do you dilute forward and reverse primers for a master mix?
I apologize for the very naive question, but I'm just starting out in a high school biology lab and I am very confused.
If I have stocks of 100μM for forward and reverse primers separately, I can dilute them to 10μM very easily. But what if I want to make a master mix? I have 1000μL of 10μM for each primer, forward and reverse (not mixed). Would I just add 500μL of my forward primer and 500μL of my reverse primer for a mix of 1000μL in  which both concentrations are still 10μM?

Would I just add 500μL of my forward primer and 500μL of my reverse
  primer for a mix of 1000μL in which both concentrations are still
  10μM?

No.  Diluting your 10μM solution in half will half the concentration.  Mixing equal parts of 10μM primer will make a master mix where each primer is 5μM
But in general, primers are added to these reactions in great excess, so 5μM of primer might be okay.

The question also states that they have 100 uM stocks of each primer. In that case, they can make their master mix (with 10 uM of each primer) using those stocks.


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Will amylase inhibitors affect the colorigenic reaction between starch and iodine?
I'm doing an experiment for my IB bio EE involving colorimetry. I'm not experienced at all with colorimetry, so I'm having some trouble planning it. The experiment is on enzyme kinetics, and I'm testing the effect of an inhibitor on the rate of digestion of starch by alpha-amylase. Currently, my plan is just to use a starch solution with an iodine indicator, add the amylase and inhibitor and measure the change in absorbance of a certain wavelength over time with the colorimeter, however, I'm worried that when I add the amylase and the inhibitor it'll affect the colour of the solution quite a lot. Will this be an issue, and if so is there a better method I could use to control for it?
Any help would be appreciated, this is my first time actually planning an experiment and I'm finding it difficult :)

Small scale testing is part of designing bigger experiments...

Agree with @SolarMike, you could see the effect of amylase inhibitor on absorbance of starch–iodine complex before proceeding with the main experiment.

Let's work through the options.
First, the amylase could affect the color of the starch-iodine complex by direct association (rather than gradually digesting the starch).  I'd think someone would have noticed that by now, but you could test amylase + starch + iodine versus starch + iodine negative control at "time 0" to see what you think.
Next, your inhibitor could affect the color of the starch-iodine complex.  In that case, a starch + iodine negative control will look different from an inhibitor + starch + iodine negative control.
Finally, perhaps amylase binds the inhibitor in some special way that causes the whole complex to hunt out starch-iodine complex and make a strange color.  It seems unlikely, yet it's not that unlikely since depending how it works the inhibitor might manage to jam up amylase in mid-bite with the starch attached, and the protein somehow manages to mess with the wrapping of starch around iodine.  In that case you need to look at whether (amylase + inhibitor) + starch + iodine looks different from amylase + starch + iodine and/or inhibitor + starch + iodine at "time 0", then consider the color change from there.
For "time 0", you might conceivably need to extrapolate back to that time point after measuring changes after some short time intervals you can accurately measure, if the color change is rapid.

To measure your amylase activity, you will monitor the disappearance of amylase’s substrate, starch.
Starch reacts with iodine (which is yellow) to form a blue compound (Amax 620 nm). This reaction is
the basis of a colorimetric assay for amylase activity. You should be very careful about the volume of the final solutions so it won't effect your results. For example you will add inhibitor to one of them and you won't add that to the other but you should add water instead of the inhibitor so dansity won't be a problem. (final volumes should be same since starch amount at the beginning will be same) You can use some acid like HCL to stop enzyme activity so it won't change the colour and won't effect your results. After the acid stops the enzymatic reaction and the iodine
reacts with the starch to produce the blue color. Any starch that has not yet been hydrolyzed by the
 amylase will turn blue color being proportional to the amount of starch
remaining. The intensity of the blue color can be by measured by its absorbance at 620 nm. The greater the change in absorbance between a sample containing the initial
amount of starch (without enzyme) and the hydrolyzed mixture containing the enzyme, the greater the
amount of starch degraded by the enzyme, therefore the greater the activity of the enzyme being
measured. Make sure you give the enzyme enough time and heat to catalyse the reaction. Sorry for grammar errors. I hope this was helpful!
Edit:you asked if inhibitor effects the experiment. I did say it above. If you use a colorless substance to denature or inhibit the enzyme it won't effect your results. If you can't do this you may add the inhibitor after the reaction is over so it won't effect the reaction but you may eliminate possible differences in color

Welcome to [biology.se]. Please have a look at the question again: it is about whether amylase inhibitors would affect the reaction between starch and iodine in any way. Can you please [edit] your answer to address that point?


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Does pheomelanin have a useful biological function?
Melanin is a natural pigment that is categorized into two main forms, eumelanin and pheomelanin. It's well documented in the science literature that increased eumelanin levels reduces the risk of developing skin cancer, whereas it was recently reported in Nature that pheomelanin may itself be carcinogenic. So my question is:
Does pheomelanin have a useful biological function?

@leonardo not at the moment, so I've removed this statement from the question. But going by my observations, there's a greater variety of redheads and blondes here compared to regions near the equator such as Africa and Asia, so I'm assuming it's down to the higher levels of pheomelanin, lower levels of eumelanin. There is also the standard skin colour map: http://www.grida.no/graphicslib/detail/skin-colour-map-indigenous-people_8b88

You seem to have forgotten to cite the paper you mention.

How do you increase the amount of EUMELANIN but not PHEOMELANIN? Do we know if this is currently been looked at by scientists?

It is worth noting all cells that produce Eumelanin produce Pheomelanin first, so it could even be a bit of evolutionary baggage. The cell has to go through stages A-B-C even though B is no longer useful directly just because something in B triggers the change to C.

I don't think that the role of pheomelanin is known yet. To produce larger amounts of it, you need mainly two conditions: A weak (or absent) MC1R signalling and the availibility of the amino acid cysteine. If MC1R signalling is strong, the preferred way goes into the direction of eumelanin. Its known for quite a while that pheomelanin produces reactive oxygen species which either directly cause DNA damage or do it indirectly by using up the cells storage of antioxidants. You can find more details in this review as well as links to a number of original papers.

My understanding has been that it aids in sun absorption in the skin, specifically in low sunlight/overcast areas. I don't have source on hand, but that may steer you in the direction you are looking for.
Also, consider that that pheomelanin, red hair specifically, may encourage breeding success, depending on where you are.

That's my guess as well, but I can't find any references. Are you a redhead?

I'm not a red head, but if you need a source. I found an article with citations.   38 Red Hair and Redhead Facts    The article may lead you to some appropriate sources, and I'll post anything I come across.

I don't think that any use has been discovered yet for pheomelanin. I can't find sources about the claim that it increases UV absorption so I guess we can't say that yet.
I would simply guess that it's not harmful enough to warrant elimination via natural selection (in places where people are less exposed to UV, that is), and may, in certain societies, have increased sexual fitness. However even this is hard to tell, because historically red hair has been considered both an identitarian, desirable feature (as for Gauls during Roman times who would even bleach their hair with limewater to obtain reddish-blond hair) and abhorrent and cause for discrimination.
Vestiges and by-products of useful mutations may accumulate endlessly if their impact on overall fitness is marginal. In this case the useful effect would be the inhibition of eumelanin the skin, which is desirable in low lit/cold climates. This may in turn cause greater pheomelanin production and allows the normally concealed pheomelanic phenotype to show.

Pheomelanin has it normal function in transforming radiant heat to energy but can't protect its host from ultra violet rays from the sun as UV will destroy melanin cells.

ahh, interesting! Do you have a reference for this?


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How much carbon does a forest contain?
Trees in a forest, alongside the fungi, animals and plants that live there in, capture atmospheric carbon dioxide and store it inside their tissues and in the soil as humus.
It is possible to estimate the amount of carbon stored in some portion (say 100 square meters) of a climax forest?
This should depend on the specific biome, for example Eucaliptus and Pine forests seem to contain less organic matter than broad-leaved tree forests in temperate regions.
PS: A slightly more speculative aspect of this topic is to evaluate weather reforestation can contribute to lower the amount of carbon dioxide in the atmosphere.

There are some well established methods for this. For example, estimating forest C stocks and fluxes is done for national C accounts which are used by the Intergovernmental Panel on Climate Change (IPCC) for global reporting.
The usual method is to measure the diameter of all trees in a defined area (a plot, e.g. like you suggest of 100 square meters), then use allometric equations to estimate either the volume or biomass of each tree, then convert this to C (which is simply 50% of the total dry biomass). This gives you the C stock of all trees in your plot, which can then be scaled up to the entire forest if you wish (you would need lots of plots to make a credible estimate).
Allometric equations vary between tree species due to their differences in shape and wood density. However the general proportion of C varies little between species, it is always close to 50% of biomass. It is better to use allometric equations specific to the tree species you are measuring but if none are available you can use a generic equations:

For live tree biomass, diameters of a sample of trees are measured and converted to biomass and carbon estimates using allometric biomass regression equations. Such equations exist for many forest types; some are species-specific, whereas others, particularly in the tropics, are more generic in nature (e.g., Alves et al., 1997; Brown, 1997; Schroeder et al., 1997). Cutting and weighing a sufficient number of trees to represent the size and species distribution in a forest to generate local allometric regression equations with high precision, particularly in complex tropical forests, is extremely time-consuming and costly and may be beyond the means of most projects. The advantage of using generic equations, stratified by ecological zones (e.g., dry, moist, and wet; see Brown, 1997), is that they tend to be based on a larger number of trees (Brown, 1997) and span a larger range of diameters; these factors increase the precision of the equations. A disadvantage is that the generic equations may not accurately reflect the true biomass of trees in the project.

If you already have wood volume data, which is often available for production forests, you can use a Biomass Expansion Factor to convert volume to biomass, and then to C.
Another method is to use lidar (laser scanning) to generate a 3D model of the trees in your plot and then calculate their volume -> biomass -> C.
Since trees are the easiest component of the forest to measure, there are methods to estimate the other C pools (litter, soil, roots, dead wood) because it is generally too difficult and time consuming to measure these. E.g. dead wood is generally 5-40% the C of live trees, but ideally you would use an estimate based on a  similar forest type.
You specifically mention climax forest - though the method is the same for all forests it is likely to be less accurate for climax forests because they tend to be more variable in tree size and most allometrics have been developed for smaller trees (e.g. in plantations) and are usually not as accurate for larger trees. To improve accuracy you would need to cut down some trees and measure all of their components to derive your own allometrics.
BTW eucalypt and conifer forests in temperate regions have the highest C stocks per unit of land area of any forests.

Thanks! I would discard the last sentence on Eucalyptus. The paper seems to refer to a single species, the E. regnans  (which can be as massive as some Sequoia, and has a lot of wood). Many Eucalyptus forests do not favour undergrowth and burn easily/regularly, thus store less biomass than others.

My comment was in response to your statement: "Eucaliptus [sic] and Pine forests seem to contain less organic matter than broad-leaved tree forests in temperate regions." This may be true as an average or total for all temperate eucalypt and pine forests, though I don't have those figures. Nevertheless, it is a relevant fact that the most carbon dense forests are certain temperate eucalypt and conifer forests.

This question is very broad and this would require a lot of calculations, I will focus on the carbon dioxide in the trees, hopefully some other users can add details in respect to the organism and their carbon dioxide storage.
I found an article(R.Jandl et al.,(2007)) which did some calculations, this site does a really good job in explaning these calculations so I will cite that:

As trees photosynthesise they use carbon dioxide (CO2) from the
  atmosphere with water from rain or irrigation and nutrients from the
  soil to form carbohydrates, which make up the tree’s biomass, but how
  much carbon is made by a tree in this process? Researchers at
  Ecometrica have worked it out! The amount of carbon stored by a tree
  depends on its size, which in turn is influenced by factors, such as
  species, local environmental conditions and the way it is managed. In
  an attempt to find a simple answer to this question, researchers at
  Ecometrica have broken down approximately how much carbon is stored in
  each element of a typical tree (the branches, the leaves, the stem and
  the roots) by percentage for a quick and simple calculation.
They used standard forestry practices to estimate the amount of carbon
  contained within the stem, branches, roots and leaves of a mature
  sycamore (Acer pseudoplatanus), found near their office in Edinburgh,
  with a diameter of 52 cm and a stem height of 12 m.
First they measured the various parts of the tree and, from these
  measurements; they calculated their volumes, which were then used to
  calculate the biomass (mass of living matter in plant tissues):  The
  radius of the stem at 1.3 m above the ground (r1 = 26 cm) and at the
  top of the stem (r2 = 20 cm), and The height of the stem (h = 12 m)
  They used known relationships between the biomass of the stem and the
  biomass of roots, branches and leaves to estimate the total biomass of
  the tree: Stem volume was estimated using the equation for the volume
  of a truncated cone:

which gave an estimated stem volume of 2.0 m3 The wood density of sycamore is approximately 620 kg/m3, so using the equation:
Mass = Density × Volume  the stem biomass was 1243 kg, or
  approximately 1.2 tonnes  The biomass of the roots, branches and
  leaves of a sycamore tree are known to be around 26%, 11%, and 1% of
  the total biomass, respectively. These proportions were used to
  estimate the total biomass of the tree without having to dig up roots
  or cut down branches.  In summary, the sycamore tree had a stem with a
  biomass of 1.2 tonnes, roots of 0.5 tonnes, branches of 0.2 tonnes and
  leaves of 0.02 tonnes, giving a total biomass of 2 tonnes.
The carbon content of woody matter (stem, branches and roots) and that
  of leaves is approximately 50% of  their biomass. Multiplying the
  total biomass by the proportion of that biomass which is carbon gives
  an estimate of the total amount of carbon stored in the tree, which in
  this case was exactly 1 tonne! Each tonne of carbon equals to
  approximately 3.67 tonnes of CO2.

Then the further calulations really depends on the density of the trees in the forest (if you know the exact number let me know). There are a lot of atricles which gave some numbers the range is based on these articles somewhere around 30-400(and some even 1200 trees). I used the data from this article, which measured the optimal density for growing. I assumed that the climax forest is just naturally so I just picked 400 trees per acre (this is just guessing offcourse and I just generalized this for all trees, but I saw 400 in an other article):

Then we can just do some simple math for the carbon dioxide for the trees:

total CO2 for the trees = 400 x 3.67 = 1468 tonnes

Let me know if someone can find the tree density of a climax forest

Great answer! I'd be happy to up vote, but I can't do that yet. I was thinking of an alternative approach, involving the notion of biomass. The more the biomass in a forest, or crop filed, the more the carbon stored. Probably, it is possible to estimate the carbon stored in a  Kg of generic biomass, and then check the how much biomass a forest has!

In a climax forest the stand density (number of trees) would be much less than 400 trees/acre. The trees would be larger though, with much more than 1 tonne of C each. Total aboveground C in trees would likely be 300-500 t/ha (1100-1835 t CO2). A hectare is 2.47 acres, so around 450-750 t CO2 per acre. Plus more for understorey plants, dead wood, roots, soil, etc.

It is certainly possible to make an estimate, and other answer have already elaborated on the methods used. As to the underlying question, how much this contributes to store carbon: Lewis et al have published a study (in Nature) estimating the amount of carbon stored over long time periods in African tropical forests. They combined this with other publications and estimated that 1.3 Pg C yr-1 is stored in the tropical forests of our planet.


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Plating 96-well bacterial transformations
I need to clone 96 different inserts into the same backbone in arrayed format. I am planning to perform Gibson assembly reactions in a 96 well plate, and then do chemical transformations in a 96 well format.
However, I am not sure how to plate 96 well transformations in the most efficient way. I could just plate them one by one onto petri dishes, dividing each petri dish into 4 sectors. However, I suppose there could be a better way of doing it. Any suggestions?

You could plate them in 6-well or 12-well plates, so you don't have to juggle all the Petri dishes.

If you want to plate them as usual you can use glass beads instead of a spreader to spread them.

I think your approach very much depends on how essential it is that you get discrete single colonies, and if this is a one off or a regular process. To preface, similar to what Maximilian Press said, using a single welled plate like a nunc omnitray would be your best bet.
The main consideration you need to take into account for high-throughput transformations is the transformation efficiency (TrE). Naturally in a big petri dish, 100 µL of a single 1:10 or 1:100 dilution of your recovery culture can bring out nice discrete colonies. When you start reducing surface area and volumes, the chance of discrete colonies obviously diminish.
The best way I've found is to do a test transformation using the backbone of your choice and gauge the average TrE over a few runs to hone in on a 'one size fits all' dilution/volume, then apply that to your full high-throughput run. Assuming your inserts are of similar size, you should see a relatively similar TrE between them (bar edge cases). Another option would be set up three 96-well plates and do three separate dilutions, 1:10, 1:100 and 1:200 at 100 µL a pop so you at least get some fudge factor.
In terms of the chemical transformation itself, doing them in a 96 well plate format will guarantee lower TrE simply due to the characteristics of the plate itself. Thicker plastic and sub-optimal heating (if you aren't using a thermocycler and PCR plate) requires a slightly longer heat shock, and recovery in media requires much more vigorous shaking due to the mixing characteristics within the wells.
If you don't care about plating out in 96 well plates, like Maximillian Press said, use a single welled plate and format it in a 96/48 well fashion. Will save a massive headache when pouring. Just make sure you leave it to dry to prevent any running culture.
In terms of actually growing the colonies in a 96 well plate, if you're making the agar yourself I'd recommend adding an extra 0.5% agarose to thicken it up (I doubt you'll be using one, but make sure your plate is non-tissue culture treated). Wait for it to cool just above its solidifying temperature (around 45 °C in my experience) and then transfer out using a pipette or low volume serological pipette to reduce the meniscus formation. If the agar forms a high angled meniscus, your colonies may form smears and will not be discrete. Leave the plate out to dry as well, colonies may also smear from the agar being moist.

TBC, I was talking about plating on a flat plate, not 96-wells! It has the same shape and size as a 96-well plate but is otherwise a flat bottomed plate. The similar format makes it easier to organize and avoid errors, but pouring 96 wells sounds like a huge headache.

Ahh yes I was agreeing with you, just mix up in what I was calling them!  I agree, using actual 96 wells is a real pain.

gotcha, cool, seems like we're on the same page.

I would suggest pouring solid media selection plates of something like this, that have the same shape and size as the 96-well plate. You can then directly spot your transformations from their wells onto this plate, which has a similar format. You will not be able to pick colonies per se (you will just have a confluent spot of cells that can grow on the selection), but if you are just interested in the constructs it would save a lot of labor and potential confusion.

An alternative to the confluent spot could be to do spot plating from dilutions as it is done in survival experiment. This would allow the recovery of individual colonies.

@GaelC a good point, though I worry about transformation efficiency being high enough to reliably recover colonies from dilutions. Maybe do a 1:10 dilution series and plate each dilution, to account for variation between reactions?

Yes, for the first time I would do that and see if 2 or 3 dilutions points could be enough for the next time.


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Can anyone identify this bug, it's got six legs
Hi I live in the midwest (Wisconsin), these have recently started appearing all over my basement. Anyone know what they might be? They are about 2.5 mm long and they are quite active. I don't keep any food items in the basement they they would be able to fed on.

Please indicate size, describe behavior, and comment about any stored food in your basement or in nearby places of your home.

RelatedTiny dark-colored bug ID in Amman-Jordan

I'd look hard through your possible food storage areas; they may not be infesting the sorts of foods you'd expect, I've definitely seen some relatives here in WI. See for examples https://extension.umn.edu/product-and-houseplant-pests/pantry-pests-insects-found-stored-food#beetles-55711 https://hortnews.extension.iastate.edu/red-flour-beetle https://hort.extension.wisc.edu/articles/sawtoothed-grain-beetle/ I'm not certain which exactly you have, I'm guessing @theforestecologist can either provide a more definitive answer here or mark as a duplicate of the related ID.


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Can a trait be too successful? Wouldn't overwhelmingly successful traits limit variability, which is one of the requirements of NS?
Can a trait be too successful?  Wouldn't an overwhelmingly successful trait soon limit the gene pool, and if so, how would the process of natural selection react to that?  If an individual is born with a trait that allows it to reproduce unimpeded, and it passes that trait on to its offspring, wouldn't the variability be quickly reduced in the population?  And if so, then would NS limit the trait so that it isn't overwhelming?
And thanks to everyone who contributed here, you have been very helpful and I am very grateful.

Welcome to Biology.SE! Please clarify and research your question. In particular, why do you think that a "overwhelmingly successful trait" would limit the "gene pool"? ——— Please also take the [tour] and then go through the [help] pages starting with [Ask] questions effectively on this site and delete or [edit] your question accordingly. Thanks!

I have found that when learning about a new area starting with a relatively accessible and reliable source like Khan Academy is very helpful. Wikipedia is also generally a good starting point and you can then check their references. Online platforms called MOOCs offer free (or very low cost) courses on a wide variety of subjects — two I am familiar with are Coursera and edX. Finally, textbooks with a good level of detail are also freely available online e.g. from NCBI.

Finally, I encourage you to check out some of the online resources available for learning more about evolution. For example, this a useful introduction to evolutionary theory from UC Berkeley. This will give you a better understanding of evolution and perhaps allow you to frame your question more clearly.

This information is very helpful and I am looking forward to using these resources to find the answer to my question.

I think you could expand the question to be more clear. See if I understood correctly: Suppose there is a given individual A in a population with a very adaptive trait T. Presumably this individual would generate more descendants, and its descendants would also reproduce at a higher rate, so that in the long run a significant fraction of the population would acquire trait T. But this seems to imply that a significant fraction of the population would descend from A and therefore a lower genetic variation.. Is this your question? If so I suggest looking for genetic drift and related things

Yes, in neutral theory of evolution this is called fixation - when all the alleles (for a given locus) become identical. Note that it may happen, even if the gene does not have a striong selective advantage - due to the random effects.
If you are looking for the basic background in population genetics, Gellespie's book is a good starting point.


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Impact of Terminator Efficiency in Genetic Constructs
I was looking at the iGEM Bioregistry of Terminator parts which offer varying degrees of termination efficiency. I am wondering why studies into combinatorial synthesis of genetic circuits for metabolic engineering include terminators as a variable component within their designs, such as this paper by Woodruff et al., 2017? This must come at great complexity cost to overall library size, where I would of assumed one would take the most efficient terminator and fix it within the design. How could terminator strength or sequence effect optimisation of a metabolic pathway in an impactful manner such that this extra complexity becomes worthwhile?

My guess is that they used different terminators to ensure sequence variability. Cloning several copies of an identical sequence into a plasmid can lead to spontaneous loop-outs, causing the additional copies to get lost over time. And for why less efficient terminators are kept in Parts Registry: why would you remove well-characterized parts and thereby lose potentially valuable data? You can only appreciate the efficiency of one part by comparing it to another.

@gaspanic Interesting, thank you for your thoughts!

To add to gaspanic's answer, not all terminators work equally well under all conditions. So if terminators weren't characterized in the organism or tissue you are working in the efficiency might be different. Additionally, though rarely people might be looking to design with a leaky terminator mimicking bacterial use of leaky terminators to use one DNA sequence to create two mRNAs of different length.
[Edit] Sources for further reading
Example of terminator efficacy differences between different species
Example of terminator differences between tissues
Terminator Attenuation - stopping of termination only sometimes
Leaky stop codons as a molecular tool

Your answer may or may not be correct, but we have no way of knowing because it is just an assertion, with no supporting evidence. Please read the Help on Answering Questions.

Sorry, I thought this was widely accepted enough that it didn't need specific references for the assertions.

@PlantPromoter This is very true that part behaviour can vary not only strain to strain but system to system (metabolically engineered system). I guess if you have the ability to implement such complexity nothing is to be lost. I don't think in this case of Woodruff et al., that readthrough would create any benefit to their system because all transcriptional units have their own promoters.


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What is the most complex biological organism (or precursors) that we have been able to synthesize from raw materials?
In the Miller–Urey experiment they produced several amino acids.  I'm not sure if there were other similar experiments that got further.
http://www.jcvi.org/cms/press/press-releases/full-text/article/first-self-replicating-synthetic-bacterial-cell-constructed-by-j-craig-venter-institute-researcher/ is a reference to making a cell, but they really injected an engineered genome into an existing cell (which is pretty amazing!)
But I'm interested in work to make completely synthetic life with no preexisting biological components.

Since you're discussing the boundary between living and non-living matter, can you provide a concrete definition for what you consider life for the purpose of this question?

a self replicating cell - but I'm open to other definitions if they exist.  And I'm also not looking only for creation of complete life - just the furthest we've gotten.  Lets imagine is the goal to make (even in lab conditions) a single self-replicating bacteria from raw materials, how far along are we?  1%, 5% 70%, etc.  I'm interested in what we've made until now and where the next steps are to go further.

This might interest you: http://biology.stackexchange.com/questions/19316/first-rna-polymerase-mrna/19737#19737

Modern bacteria are extremely complex (they've been evolving for about 4 billion years) and probably look very little like the first organisms on earth. We're not even sure if early life used DNA. Whatever we can produce in a lab from no pre-existing components, is bound to be hard to recognize as life as if it had evolved on an entirely different planet.

I think the simple answer is that no one has come anywhere close to doing what you ask for, under any commonly accepted definition of "life". Jack Szostak's work  (http://molbio.mgh.harvard.edu/szostakweb/) is a good place to start researching this.

@LeonAvery - if you want to post your comment as an answer, I can select it as no one else has offered anything better.


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Where can you find the quantities of each amino acid of a particular protein or food?
Taking a potato as an example. If I wanted to know how much µg or % of each of it's amino acids there are in 1 gram of pure potato protein, where can I find this information?
Is there a freely accessible database online with these statistics? Or, one that contains something for each protein from which this could be derived? (and if so how?)

you mention as an example a "potato" - I put a link to the USDA database that lists a number of foods and protein sources and the known nutritional info

Both excellent answers so far, I'm a bit short on time so I haven't figured out yet which answer is effectively best to accept, though I've been looking into both, but rest assured there will accept properly and upvote in a day or 2 max.
P.S.: Should I expect much fluctuation in bio-availability of the amino acids for each protein? Or is bio-availability more of a thing to worry about for other compounds and not at all for proteins and amino acids?

The bioavailability shouldn't fluctuate much for the protein, which should be fairly constant for each of the individual amino acids - I can't comment what the bioavailability is for the nutrients other than the macro nutrients (i.e. Carbohydrates, fats, amino acids)

There's a fantastic database available from the United States Department of Agriculture that includes almost 9,000 common foods, including their nutritional information. This database is searchable and available from the USDA Agricultural Research Service. Here is a link for the online searchable database.
Within the database you are able to search for a number of parameters; including, food composition, water content, individual amino acid content, fatty acid and total fat content, vitamins, minerals, phytonutrients, and more! One of the nice things is that you can ask it to rank the foods in order of nutrient composition. For example, ranking all the foods in the database in order of protein content or a specific amino acid (or whatever else you're interested in).
It's a great database, especially for researchers that need a database of trusted nutrient data.

Within the database you can search by nutrient or by food, whichever you are curious about. For example, as you mention in the question, I entered potato and in the second screen shot below you can see the listing of all the nutrients in the potato. These are listed as % weight per 100g of potatoes - with or without the skin.

To get to the amino acid content breakdown, click on Full Report (All Nutrients) at the top.

Which question to upvote between you and Samuel is a brain breaker. The original question was per gram protein, I assume you changed my question to account for foods that contain additional lose amino acids. However I effectively needed per gram pure protein from the food rather than per g of food. Anyhow I've upvoted your question.

@Happy the USDA database gives you both the total amount of protein and the total amounts of amino acids (as well as all the other nutrients) - you can divide one by the other to get amino acid per gram of protein. Now I think I'm confused by what you're saying... ?

Yeah in Samuel's case an additional calculation seems to be also needed so I think I'll just settle on your answer as accepted one.

Original link no longer works. New site seems to be https://fdc.nal.usda.gov/

Go to WolframAlpha and enter 1g potato in the input field:

Then scroll down in the results to Protein and amino acids and click the more button in the top right hand corner to get the following list of amino acids with their compositional values for one gram potato:

you need to know the protein sequence to be able to do this.
Paste the protein sequence into this tool and press 'Compute Parameters'. This will give the percentage amino acid composition of the protein.
http://web.expasy.org/protparam/
Otherwise if you mean 'generic protein from potato' then as a quick google reveals this
http://journals.cambridge.org/download.php?file=%2FBJN%2FBJN12_02%2FS0007114558000271a.pdf&code=829c6412f21656bf0e528452fc6608c7
I am not sure there is a database of amino acid composition of foods
EDIT
Actually I found this resource, might warrant further inspection
http://www.fao.org/docrep/005/AC854T/AC854T00.htm

Good answer, I've upvoted, sadly for example for almonds the amino-acids are expressed only with A: mg/g total nitrogen, which I don't understand how to interpret as it differs wildly from B: mg/100 g food. So I don't have usable data for almonds. Oh and thank you for the cambridge article, interesting to consult it in comparison to the database.


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How are targets formed for axon growth cones (CNS)?
Axons have growth cones which find a route to their target using multiple methods (guidepost cells, attraction to target, etc...). My question is, what is the process that actually forms the target? For some given Axon A1 of Neuron A, why does it skip Neuron B, and go straight for Neuron C? How is a target formed on neuron C for Neuron A to connect to? Are there are any articles/readings on this topic? There is also the process of following the path of a pioneering axon. Does following a path of a pioneering axon take precedence over other forms of signaling? Does distance play a factor in choosing a target (Eg: Axon growth cone prefers closer target rather than distant target?)


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How can rooted and unrooted (phylogenetic) trees be distinguished from one another
I understand that rooted and unrooted phylogenetic trees are similar in structure
But how can they be easily identified as one or the other?
Is it simply based on the presence or lack of a named root (the root is identified as a particular ancestor) or outgroup?

At which "stage" do you want to identify the tree? When it's a file (Newick), or when it's a drawn figure?

I'd say a drawn figure?
Like something you would find on a google search for phylogenetic tree (http://images.google.com/search?tbm=isch&q=phylogenetic%20tree)

In case of a drawn figure, either look for the a short stem to the left, or for an explicitly labelled outgroup. Trees derived from phylogenetic analyses are normally unrooted by default, you need to root them by either making an arbitrary (but hopefully informed) decision or by adding an outgroup.

So when you say root it by making an arbitrary decision, does that mean labeling a particular ancestral individual as a root?
Also, does this consider midpoint rooting?

Midpoint root is certainly one of the ways you can choose to root your tree. IMHO rooting a tree of species or proteins without an outgroup is done by adding an extra root node.

And just to clarify, how would the 'extra root node' appear on a diagram?
As just a labeled point from where the rest of the tree branches off from?

Usually, there's a short stem going in a different direction than all the other leaves. But a labelled point is also fine, I guess


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Richard Goldschmidt's “systemic mutations”?
[TLDR] I dont understand the fault people had/have with Richard Goldscmidt's concept of systemic mutations [/TLDR]
As a preface, I am a undergrad biology student, so sorry if this is a "Stupid question".
How does Richard Goldschmidt's concept of systemic mutations, reject the classical gene concept, why were biologists so against it?
From what I  understand he used the concept of systemic mutations and developmental macromutations in his lectures to explain how macromutations could cause speciation(macroevolution?). The results of the developmental macromutations, if I understand him correctly would be then the Hopeful Monsters?
And from what I have read on a timeline of his work, systemic mutations describe the "large rearrangments of the chromosome", with a new arrangement meaning a new phenotype?
How is that wrong though? Do chromosomes not have rearrangements via cross over events during meiosis or via transposable elements? Or is he referring to something else, since I understand at the time, meiosis and TE's were not understood well.
The source i was reading from was:
(Richard Goldschmidt: hopeful monsters and other 'heresies' by Michael R. Dietrich) Available from http://www.dartmouth.edu/~dietrich/NRG2003.pdf

His notion of systemic mutations involved the postulation of massive chromosomal rearrangements (not mere recombination/crossing-over) as mediators of speciation in one-step. While whole genome duplications have been shown to induce speciation (see, for example, cryptic speciation in Hyla versicolor) they are not large scale rearrangements as he suggested. Additionally, while there is evidence for multiple large scale chromosomal arrangements at once in cancers (Chromothripsis) , there is no evidence of a similar process resulting in immediate reproductive isolation.


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Was site-directed mutagenesis used to study the effects of sickle-cell anemia?
The title says it all. I am inquiring whether or not site-directed mutagenesis was used to study the effects of sickle-cell anemia/figuring out that specific amino acid mutation was the problem.

Why mutagenesis? You can simply genotype and see what the mutation is.

Mutagenesis was brought up in my lecture and how it can be used to see the affects of single amino acid changes. I asked the professor whether or not they could apply it to sickle cell anemia and there wasn't a clear answer.

The cause of sickle-cell anemia to be a molecular defect was discovered in 1949 by Linus Pauling. In his experiment hemoglobin molecules were broken down to smaller fragments, which were then subjected to electrophoresis and chromatography. The results of these measurements were compared between healthy and affected samples, allowing the identification of a mutated fragment.
The amino acids in the mutated fragment were analyzed in 1956 by Vernon Ingram, identifying the single mutation. The process in a bit more detail is described on this webpage.
Site-directed mutagenesis is used primarily in research aimed at finding a gene therapy based treatment for sickle-cell disease, such an attempt is described in this publication.
Also, the technique was used on recombinant hemoglobin to create and study some properties of naturally not occurring mutations. [Link]


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Is genome recurrence possible like in Jupiter Ascending?
I have watched Jupiter Ascending many times now and I was wondering if genome recurrence, like what happened to Jupiter Jones, is possible.
Is it possible to find two unrelated people, possibly several generations apart, that cannot be distinguished between by DNA sequencing?
Taking the example of Jupiter Ascending: in the movie Seraphi Abrasax they show the "genetic recurrence" of Jupiter Jones, which means the same sequence of DNA has repeated after several thousand years from Seraphi to Jupiter.

Could you expand on "genetic code recurrence like what happened to Jupiter Jones". This is new to me. It sounds like you are asking "is it possible for unrelated people to be genetically identical?" Technically possible. Statistically, however, nigh on impossible.

@James I'm directing you to this wikia http://jupiter-ascending.wikia.com/wiki/Recurrence for more information.

I got the gist of it. For all intents and purposes, it's impossible.

I have edited this post, especially the title, to replace "Genetic Code" by "Genome". If you do not understand the difference between the two, follow the links I have provided.

@James if you understood the question, do you think you could try to clarify it by editing the question but I don't understand the question!

@Remi.b It's fantastical and loaded by saying "is it possible" (anything is possible given enough imagination), but I've tried to edit my interpretation into the question. Richeve, if I have changed the meaning of the question, please do roll back.

Various types of genetic variation exist in natural populations. One is the single-nucleotide polymorphism (SNP), which is where different nucleotides are present at the same location in the genome of different individuals in a population (with variation present in a nontrivial number of people; the threshold is usually set arbitrarily at 1%).
Even just considering SNPs and ignoring other types of variation, what you talk about would be effectively impossible.
The human genome is about 3 billion base-pairs long. Along the whole genome, according to Chen et al 2012, between two individuals there is something like 3 million SNPs. Imagine two people flipping a coin and the result is exactly the same every time, 3 million times. Now imagine instead of heads or tails, there are 4 options.
Only 1 result of the series of 3 million 1 in 4 chances is identical to the original series.
$\frac{1}{4^{3000000}}\approx\frac{1}{\infty}$ = 0
Go ahead, try it on your calculator.
Now, there are also variations that are beyond SNPs between individuals that cause much more variation between two people. These can cause even more "unprobability". Since we already require infinite people before this event is likely to happen, I think we can just call this event impossible.

3 Gbp is 3 billion, not 3 million, so the odds are even lower. It's not = to $1/\inf$ though. Its still finite, its just vanishingly small if we are being pedantic :P

That we need infinite people does not make it mathematically impossible (as you state in a comment), but biologically (or generally, practically) impossible.

@JoeHealey I was coming at it from the point of a comparison between 2 people: of the 3 Gbp in their genomes, 3 million bp are likely to be SNPs from one person to the other. The likelihood that those SNPs are the same is an incalculably small number (at least, according to my calculator :P).

Ah I see what you meant, yeah.

It is almost impossible. Considering that the haploid Human genome is around 3 Gb (Giga-basepairs) and that ideally any of those can mutate generation after generation, the chance of having the very same sequence recurring in a population is extremely unlikely. There will be some differences.

I was thinking that your term "population" refers only to Earth, but in the movie, the population refers to their "planet farms" and they have many seeded planets that uses their own genetic code as the base to construct new sets of modified human beings. So with these large set of population it is still not possible?

Still, you have 4^3giga possibilities (roughly speaking). Even having the whole universe and billions of years it will be an "almost-zero" chance of having two identical sequences. Unless you interfere with technology, some cloning-like tech for example. But if we stick to natural reproduction, then the chances are ridiculously low.

I was trying to picture this mathematically, like if you have a finite set of bases and the DNA sequence which is long but non-continuous which means there's a limit to its length then any sequence with finite set of elements and constant length can have a possibility of repetition even if the generation is random. The chances is almost zero but not zero which means there's still a possibility. Since Earth is a closed space and considering we haven't tapped yet to cloning, or haven't introduce synthetic bases to our DNAs then our DNA set is still finite and thus repetition is still possible.

Mathematically possible, practically impossible. That's why my answer states "almost impossible".


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Is NADPH salt still usable after an hour at room temperature?
I accidentally left a delivery of NADPH Tetrasodium Salt (Santa Cruz Biotechnology) at room temperature for a little over half an hour.
In addition, when I opened up the package, I briefly held the bottle by the body instead of the cap (which would have prevented body heat from contacting it).
It was packed with airbags, but no ice packs, which is why I am asking this: is the powder still stable and usable?

AliceD - I rolled-back part of your edit because I think the original meaning was lost. If you disagree, feel free to go back to your version.

@canadianer - you're the expert on these matters :) I'll go for it :)

@AliceD Well I wouldn't say that ;) Actually, whether the bottle was held by the body for a second instead of the cap hardly seems relevant to the question.

@canadianer - yeah, I reckoned half an hour RT instead of -20C (dT=~45C) is a bigger deal than a second of bodyT-RT= 12C :)

Nothing significant would happen in half an hour (especially for a sealed pack of anhydrous powder). -20 is for a moderate long term storage.

Since the delivery wasn't temperature controlled, the contents will have been at ambient temperature for essentially as long as it took to get from their warehouse to your site, which could be hours or even days. Is an extra half-hour likely to be significant?

In my experience, half an hour at room temperature will make absolutely no difference.  Boehringer Mannheim (now part of Roche), who at one time supplied the best NADPH, used to recommend storage at 4o C.
By A340 callibration, NAD(P)H is typically about 85% 'pure' based on dry weight measurements, and Sigma will try to tell you that the remainder is mainly water.  That is, if you calculate a concentration based on the dry weight, the A340 will be about 85% what you expect.
Even if some water is absorbed from the atmosphere, this will make little difference as you are probably going to calculate the concentration based on
the A340.
As User 243 has pointed out, if you are concerned that the NAD(P)H is degraded or oxidized, then an A340
measurement will probably set things right.
If you are concerned that all the A340  absorbing material is not NAD(P)H, or that a significant proportion is in the α-form (Oppenheimer & Kaplan, 1975) or is otherwise enzymically inactive, the best thing is an enzymic calibration.
This is done by adding limiting amounts of NAD(P)H to an assay system containing excess substrate and calibrating enzyme, and measuring the total decrease in absorbance (Racker, 1957).
A popular calibrating enzyme for NADH is aldehyde dehydrogenase (EC 1.2.1.3), as the reaction is practically irreversible, and it makes it easy to 'drive' the calibrating reaction to completing (and you don't have to worry about equilibria).
As I said in a previous post, NAD(P)H is UNSTABLE IN ACID (Oppenheimer and Kaplan, 1974). To put it crudely, 'the head (nicotinamide moiety) falls off', causing a bleaching of the absorbance at 340nm.  Making 100mM acetate, pH 5, 100 micromolar in NAD(P)H produces a very significant decrease in A340 absorbance with time (which may be very easily monitored in a spectrophotometer).
Just because all the A340 material is due to NAD(P)H, this does not necessarily mean that the commercial preparation is pure.  In fact, far from it.  Dalziel (1963) showed that commercial preparations of NADH then available contained a competitive inhibitor of many dehydrogenases, most probably adenosine diphosphate ribose.
Accepting Dalziel's value of 17 600 M-1cm-1 for the extinction coefficient for NADH at 260nm, and assuming that NADPH has an identical value, then highly purified samples of NAD(P)H should have an A260/A340 ratio of about 2.83, and values higher than this may indicate the presence of impurities that absorb at 260nm, but not at 340nm. ADP-ribose is a prime example, but NAD(P)+ is another possibility.
I notice that in one of the links given by AliceD, Sigma quote a A260/A340 ratio of 2.32. So is the higher value due to the presence of residual NAD(P)+, or something else (a breakdown product?), or both?
You may need to use chromatography, typically on an anion-exchange resin, to obtain NAD(P)H free of impurities.
References
Dalziel, K. (1963). The purification of nicotinamide adenine dinucleotide and the kinetic effects of nucleotide impurities.  J. Biol. Chem. 238, 1538 - 1543.  [pdf]
Oppenheimer, N. J. & Kaplan, N. O. (1974). Structure of the primary acid rearrangement product of reduced nicotinamide adenine dinucleotide (NADH). Biochemistry 13, 4675 - 4685.[pubmed]
Oppenheimer, N. J. & Kaplan, N. O. (1975). The alpha, beta-epimerization of reduced nicotinamide adenine dinucleotide.  Arch. Biochem. Biophys.166,  526 - 535.
Racker, E. (1949). Aldehyde dehydrogenase, a diphosphopyridine nucleotide-linked enzyme.  J. Biol. Chem. 177, 883 - 892. [pdf]
Racker, E. (1957). Spectrophotometric enzymatic methods for the determination of aldehydes and ketoaldehydes.  Methods Enzymol. III, 293 - 296.
Other References
Ciotti, M. M. & Kaplan, N. O. (1957). Procedures for determination of pyridine nucleotides.  Methods Enzymol. III, 890 - 899
Horecker, B. L. & Kornberg, A. (1948). The extinction coefficients of the reduced band of pyridine nucleotides.  J. Biol. Chem. 175, 385 - 390. [The definitive determination of the extinction coefficient at 340nm] [pdf]
Kornberg, A. & Pricer, W. E. (1950). On the structure of triphosphopyridine nucleotide.  J. Biol. Chem. 186, 557 - 567.
[This paper shows that the 'extra' phosphate in NADP(H) is attached to the  2' position of a ribose.  In Coenzyme A, of course, the phosphate attached to the ribose is on the 3' position. [pdf]
Pullman, M. E., San Pietro, A. & Colowick, S. P. (1954). On the structure of reduced diphosphopyridine nucleotide.  J. Biol. Chem. 206, 129 - 141 [pdf]

You could measure OD at 340nm. If OD340 is much lower than expected, NADPH is oxidized and does not have much biochemical activity.
http://www.bmglabtech.com/media/35216/1043734.pdf

I think when they sent it to you without dry ice, it is probably OK to store it at room temperature.
Sigma seems to advise on their NADPH to store dehydrated NADPH at room temperature, while they advise to store the hydrated forms at -20oC. Likely the presence or absence of molecular water in the material is crucial for its storage.
disclaimer: I am not an organic chemist and when you are doing crucial experiments with your material and you doubt its quality, I'd recommend to chuck this batch away and order new. Likely, a single day of futile experimentation in the lab due to bad reagents is more expensive than the purchase of another jar of NADPH.

@TomD - the water in NADPH-hydrate is not considered part of the molecule. It is part of the crystal. Therefore it doesn't show up in the formulas. The stability difference is because NADPH reacts with water I guess. The dehydrated form is called the anhydrate. I just talked to a chemist about this :) And I linked the question to him too. btw: I +1'd your answer. Great stuff.

@TomD - quite frankly I don't know, and my friend did neither. The only clue I got was the fact that the hydrate salts need storage at -20oC, the anhydrate form not. I'm not a chemist (as posed in the disclaimer :-)


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Agarose gel Ladder smear
We are having trouble with agarose gel electrophoresis. It used to work a couple of months ago but now the ladder always look smeared. We switched the components (1x TBE, 100bp ladder, different type of agarose and voltage) and equipment. We also tried 1.2%, 1%, 1.5% and 2% agarose.
I would appreciate it if someone can help us.

Was the alternate ladder new, the agarose? If all your components are old or being improperly stored or handled and thus degraded, you will get smearing.

Are you sure that your TBE is at the right dilution? And that you're using TBE to make the gel and also as the buffer for running?  That sort of wavy line looks like what you might get if you'd made the gel with water instead of buffer, or run it with water instead of buffer.  You could also try using TAE to make/run the gel as a sanity check.

I second the possibility of gels cast with water instead of TBE buffer. It is a classic mistake that many people make the first time in the lab. Another classic would be making gels from a 10x TBE stock and using 1x TBE for the run.

Try going even higher with the agarose concentration, as it looks like you have decent separation of the small bands and poor separation of the large bands. Also try a fresh batch of agarose.
I would suggest trying 2-4% in 0.5% steps
If you are running 100bp or smaller fragments, you should in the range of 4% agarose.
Here is a nice example of agarose ranges and minimum difference of PCR products that can be resolved. https://www.qiagen.com/mx/resources/faq?id=1b23a0b3-32b4-4c1b-bd84-49b4b6a58311&lang=en