text:
Antibacterial hand soaps (and related products); what are they good for?
From Wikipedia:

Household use of antibacterials in soaps and other products, although not clearly contributing to resistance, is also discouraged (as not being effective at infection control).

Following the links led me to this paper:  Larson EL, Lin SX, Gomez-Pichardo C, Della-Latta P.,  Effect of antibacterial home cleaning and handwashing products on infectious disease symptoms: a randomized, double-blind trial.  Ann Intern Med. 2004 Mar 2;140(5):321-9.
They conclude:

The tested antibacterial products did not reduce the risk for symptoms of viral infectious diseases in households that included essentially healthy persons. This does not preclude the potential contribution of these products to reducing symptoms of bacterial diseases in the home.

Upon reading this, my thoughts were:  Why would one test whether antibacterial products have an effect on viral infections diseases?  Since the authors used a control group (those whose soaps didn't have an antibacterial agent), they are only testing for the effect of the antibiotic.
This leads me to the main question:

Question:  Which common diseases would likely be more affected by antibacterial hand soaps (and other antibacterial products) than their non-antibacterial counterpart?

psychosomatic illnesses?  Soap is a pretty potent anti viral and anti bacterial all by itself.  Hospitals and doctors use Purelle or other alcohol based gels to keep their hands clean.  The weak antibiotics in antibiotic soaps like triclosan are probably encouraging the formation of antibiotic resistance.

This may be related to this question on Skeptics SE:  Does regular handwashing with antibacterial soap prevent disease?  Since it is closely related, I will copy my answer from there to here:
See this related answer.
To answer your specific questions:

Has something changed?

Well, we have access to these products now, whereas we didn't before.

Does this sanitizing really make people healthier?

That is a subject of debate.  While it can prevent rapid spread of some things, keep in mind that anti-bacterial soaps should only have specific uses.
The CDC has concerns about the prevalent use of these products.

Scientists are concerned that the antibacterial agents will select bacteria resistant to them and cross-resistant to antibiotics. Moreover, if they alter a person's microflora, they may negatively affect the normal maturation of the T helper cell response of the immune system to commensal flora antigens; this change could lead to a greater chance of allergies in children. As with antibiotics, prudent use of these products is urged. Their designated purpose is to protect vulnerable patients.

Also, there is some concern as outlined by the hygiene hypothesis (WP).  However, that is mostly associated with childhood immunities and allergies as opposed to the effects on adults.

“The natural immune system does not have as much to do as it did 50 years ago because we’ve increased our efforts to protect our children from dirt and germs,” says McMorris.
“Allergies are on the rise because our society has changed the way we live. As a result, people with allergies are having children with others who have allergies, which in turn creates a natural increase in the prevalence of allergies in our society.

The bottom line on this hypothesis is that more research is required.

He (Richard G. Barbers, M.D., USC professor of medicine at the Keck School) also gives credence to the hygiene hypothesis in that "we may be over-protecting kids, and their immune systems, consequently, do not become well-developed."
Barbers notes that only time and a lot more research will tell whether the hygiene hypothesis is valid, and whether, once again, parents should let their kids play in the dirt.

It is generally understood that interaction with beneficial germs is good for you.  However, normal cleaning will not destroy these germs since they reside within you (unless his germaphobia induces him to take antibiotics).  The human immune response doesn't particularly work that way.

The investigators show that "good" bacteria in the gut keep the immune system primed to more effectively fight infection from invading pathogenic bacteria. Altering the intricate dynamic between resident and foreign bacteria - via antibiotics, for example -- compromises an animal's immune response, specifically, the function of white blood cells called neutrophils.

Using too many anti-bacterial soaps has raised quite a few concerns even still.

Soaps and lotions that include antibacterial agents have no benefit over ordinary soap and water, but more research is needed to allay or substantiate concern that these substances may be leading to increased rates of antibiotic resistance. This is the conclusion of the Food and Drug Administration's Nonprescription Drugs Advisory Committee, which met last month to consider the use of these products outside of the health care setting.
"In the absence of proven benefit, there's no real reason to encourage the use of these products," said Alastair J.J. Wood, MD, committee chair and associate dean at Tennessee's Vanderbilt Medical School.

That is not to say that their use should be discontinued entirely.  Two MDs weigh in and conclude:

Opt for regular soap and water, unless you’re at a hospital or doctor’s office, where it’s best to use antibacterial soap. But if you’re out and about and antibacterial soap is the only kind around, it’s fine to wash with it—and it’s worth keeping in the kitchen to use after handling raw meat.

Hope that helps.

I thought I would attempt to answer my own question.  The only other answer currently (by Larian LeQuella), while a helpful comment, I feel it doesn't answer my original question.
To begin, we need to put the results into context.
Fact: Hand-washing using tap water alone will reduce the amount of bacteria and viruses on hands (they might not be actually killed, but no longer on your hands).
Some examples:

Ansari et al. (1989) performed an experiment where a fecal solution was placed on participants' hands and subsequently washed off using one of a range of soaps.  They found a reduction in around 83% of the human rotavirus and 90% of the E. coli.
Shojaeia et al. (2006) randomly selected 150 food handlers in Iran, and instructed them to wash and scrub their hands with sterile water.  Before the intervention, around 73% were contaminated with bacteria (primarily with Staphylococcus aureus or E. coli), and afterwards 32%.

Fact: More bacteria and viruses are removed from hands by hand washing by using some type of soap (not necessarily antibacterial).
Some examples:

Ansari et al. (1989) (op. cit.) found that certain agents (e.g. 70% isopropanol) can increase the reduction in E. coli and human rotavirus to around 98%.  However, this was not true of all possible hand-washing agents.
Mbithi et al. (1993) also conducted a experiment consisting of placing a fecal solution on participants' hands that was subsequently washed off using one of a range of soaps.  In this case, the hepatitis A virus (HAV) and poliovirus type 1 (PV) was considered.  They found that tap water gave a reduction of HAV of around 80% and PV of around 85%.  The reduction increased to around 88-92% for HAV and 90-98% for PV for most soaps tested.

[There will be zillions of additional examples to the above.]
A large survey was conducted by Aiello et al. (2007) which considered the efficacy of triclosan (one of the common active ingredients used in antibacterial hand soap) found that, overall, previous research has indicated no significant additional benefit vs. ordinary hand soap.  They write:

Soaps containing triclosan within the range of concentrations commonly
  used in the community setting (0.1%–0.45% wt/vol) were no more
  effective than plain soap at preventing infectious illness symptoms
  and reducing bacterial levels on the hands.

This is probably not the last word on the matter, e.g. Fischler et al. (2007) claimed a significant difference in the transmission and acquisition of E. coli and Shigella flexneri after using an antibacterial soap.  It is also plausible that antibacterial agents other than triclosan (where the focus has been) provide a health benefit (for example, methylchloroisothiazolinone/methylisothiazolinone).  In fact, the Aiello et al. (op. cit.) paper itself indicates a non-negligible effect from >1% triclosan.  Kimberly-Clark give a 99.9% or more bacterial killing efficacy for a range of bacteria in a 2003 in vitro experiment of their product; obtainable via Googling "kimberly-clark killing efficacy".
(Side note:  There is also a suspicion that the use of triclosan will give rise to triclosan resistance or cross-resistance to antibiotics (e.g. by Levy (2001), Aielloa and Larson (2003), Yazdankhah et al. (2006)).  Although, Cole et al. (2003) and Weber and Rutala (2006) claim otherwise in the case of antibiotic resistance.)
This leads to the original question:  Which common diseases would likely be more affected by antibacterial hand soaps (and other antibacterial products) than their non-antibacterial counterpart?  I.e. if we wanted to test the claim that a certain antibacterial hand soap has some positive effect (vs. a non-antibacterial version), what would be some common diseases that we could test for, and expect to find a non-negligible result?
I offer the following candidate for a bacterial disease whose effect should be noticeably reduced by the use of antibacterial hand soaps:

Staphylococcus aureus is a bacterial species that causes a range of noticeable illnesses and can be transferred by skin-to-skin contact and contact with contaminated objects.  We can thus reasonably expect it to be affected by an effective antibacterial hand soap.  In fact, washing hands is recommended for reducing the prevalence of S. aureus, along with zillions of other diseases (by e.g. Better health channel).  This is one of the bacteria that Kimberly-Clark give a 99.99% kill efficacy of their product in vitro.

Although, not directly related to hand-washing, Brady et al. (1990) used a 1% triclosan preparation and Zafar et al. (1995) used 0.3% triclosan to control methicillin-resistant Staphylococcus aureus.

The single additional measure of changing handwashing and bathing soap to a preparation containing 0.3% triclosan (Bacti-Stat) was associated with the immediate termination of the acute phase of the MRSA outbreak.  -- Zafar et al.

Similarly, a 1.5% triclocarban bath has been used to treat atopic detmatitis Breneman et al. (2000):

The antimicrobial soap regimen caused significantly greater
  improvement in the severity and extent of skin lesions than the
  placebo soap regimen, which correlated with reductions both in S
  aureus in patients with positive cultures at baseline and in total
  aerobic organisms.

The above results suggest that antibacterial soaps could have a beneficial effect against S. aureus (more than a non-antibacterial counterpart).  However, it is unclear as to how much of this effect implies a comparable effect for hand washing alone (and in an ordinary household setting).  Moreover, there are many different antibacterial hand soaps and other products, whose active ingredients might be at different dosage, all of which would further affect the outcomes.


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What did pangolin scales evolve from?
Were they developed anew or a heritage from fish?

Pangolin scales are certainly not retained from fish. Pangolins are within the clade Pholidota (Tree of Life page for Eutheria) and their nearest relatives are anteaters, sloths, and armadillos.
According to the University of California Museum of Paleontology, similarities between pangolin and armadillos are the results of convergent evolution. So the pangolin scales are a new feature that arose somewhere on the lineage leading to pangolins.

were they developed from hair or from skin folds?

They are keratinous scales, like hair and claws/nails. I'd suspect similar development to nails, but I'm not sure.

Their closest relatives are carnivores, not anteaters, sloths, and armadillos.  See DOI: 10.1186/1471-2148-6-93.

source: Stack Exchange

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

sort: votes


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Fetal development, gastrulation and embryonic disc
I am completely confused by the images circulating on the internet of human gastrulation.
First, lets see how it happens in deuterostomes. This image depicts the process:

(image is from Wikipedia)
From here we can conclude that blastula becomes gastrula when some of the ectoderm beecomes endoderm, the place where it goes inside becomes anus and then mesoderm is formed.
Ectoderm here is the outermost level and gastrula already has anus.
However, in this image of human gastrulatuion we see completely different things:

(image source)
The mesoderm here is the outermost layer, ectoderm is inside and the gut cavity is formed by the separation of a part of the yolk sac. There is no anus and the posterior end of the gut is blind. Also other similar images suggest that a twins may be separated already after the mesoderm was formed (thus uniplacental twins).
The lack of agreement between the images disturbs me.
Just to point out some of the differences in the depicted processes:

In first image mesoderm forms after gastrulation, on the second it forms far before gastrulation
In the first image anus forms in the process of gastrulation, in the second image anus remains blind
In the first image ectoderm is the outermost layer, while in the second picture mesodem is the outermost level that encloses all, including ectoderm.

Two suggestions: Please link to the source of the images, and please state what your actual question is.

Human embryogenesis is much more complicated than the more general and simplified picture given in your 1st figure.
For details, please study http://en.wikipedia.org/wiki/Human_embryogenesis
To point out the most distinctive features which might have led to your confusion:

Before gastrulation, the blastocyst cavity is formed.
Only the inner cell mass undergoes gastrulation.
Mesoderm forms quiet late and spreads, thereby also forming the chorion as well as the amnion.
The embryo itself is formed from a subset of all embryogenetic cells. This is shown in your 2nd figure, part C and D. Yet, tissues are still going to spread such that the final ectoderm as well as mouth and anus will be formed.

The second picture does not show human gastrulation.
Your source talks about the development of yolk sac in human embryos, and does not claim it depicts gastrulation. In human embryos gastrulation happens about 7 days after fertilization and the development of membranes, as depicted in your second picture, begins about 15 days after fertilization.
I highly recommend the part of Chapter 11 from "The Developmental Biology", which deals with early mammal embryo development, from fertilization till formation of the body axes.

How it turns that mesoderm is the outermost layer in the second picture and not in the first one?

Second picture doesn't mention mesoderm. Look closely at the titles. It's just that colour schemes are different, so one thing is blue in the first one and another in the second one.

Mesoderm is red on the second picture. It is tissue that forms heart.

Also ectoderm and endoderm are shown in direct connect to each other in the second picture while on the first one they are separated with mesoderm

The Wikipedia protostome vs. deuterostome figure depicts a "prototype" deuterostome, such as an echinoderm or an amphibian.
In amniotes (reptiles and their descendants - birds and mammals), the giant amniotic egg full of yolk led to profound changes in early embryogenesis, including the disappearance of the true blastopore. (Its analogue as organizer of gastrulation is the Hensen's node, but it is dorsal and does not form the anus.) Placental mammals have secondarily lost the amniotic egg, but a substantial part of their gastrulation movements still reflect its former presence.
Although in us the blastopore does not form the anus and does not even exist, we are still classified as deuterostomes because of our clear kinship with amphibians.

This gives a third theory - that the proto-anus is homologous to neural tube!

By your theory how neural tube is formed in amphibians then if they have blastopore which becomes anus?

What is the question?

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


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Which is more important for protein expression mRNA structure or codon optimization?
The field seems extremely divided on the debate. On one hand, artificial experiments have suggested that synonymous mutations don't correlate with gene expression but rather, the mRNA 5' structure is the most important 1. On the other hand, genome wide analysis suggests that tRNA biases are better associated with high expression 2. What other works balance out this discussion?

Coding-sequence determinants of gene expression in Escherichia coli
Translation efficiency is determined by both codon bias and folding energy
An evolutionarily conserved mechanism for controlling the efficiency of protein translation

This is an excellent question! To my knowledge, there hasn't been a definite answer yet. Recently, I did tons of research on which factors influence protein expression and you should definitely check out the following questions which I asked:

What is the criticality of the ribosome binding site relative to the start codon in prokaryotic translation?
What determines a successful protein expression in E. coli?

I answered my second question by posting a concise version of a paper from DNA2.0 (a gene synthesis company) that discusses the protein expression. It summarizes all the important factors (RBS site, codon frequency match, 5' mRNA secondary structure), but it doesn't discuss the extend to which they influence expression.
There is a nature paper from Voigt lab about RBS calculator, which takes into account only the 5' mRNA secondary structure and it doesn't discuss at all the codon-optimization. The calculator was good, but far away from perfect: there is a 50% chance of predicting an expression level within a 2-fold range of the target one.
On the other hand, an older paper from DNA2.0 argues that the mRNA is covered with ribosomes practically all the time, so the secondary structure of the mRNA should have little effect. Besides, an actively translating ribosome can break up stem-loop structures.
In my opinion, we are still far off from predicting RNA secondary structure formation, although it might be highly implicated in protein expression. From what I read, I think that it is equally crucial that there is no strong secondary structure at the 5' end of the mRNA, the RBS sequence is close to the consensus one and it it appropriately spaced upstream of the start codon, and the first 60-100 codon frequencies match the frequency of your heterologous host.

nice answer Gergana!

I spent quite some time trying to answer the same question, so I couldn't help but share my knowledge!

Consider mRNA primary structure in terms of uORFs, upstream open reading frames. These can cause ribosome pausing or slowing and even disassociation from the mRNA.
I agree that a definite answer is not known (+1). For different mRNAs, under different cellular conditions (eg, low levels of a given charged tRNA), the balance or interplay between structure- and codon usage-based control of translation could be different.


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Can genetically modified genes jump to bacteria in the eater's intestine?
The Guardian ran an article a while back talking about GM gene's jumping to bacteria in an eater's intestine. Has other research confirmed this phenomenon?

It is intestine, not stomach. Also, I love how the article says that because it can happen in certain conditions for bacteria that live in bees guts "The research - which has yet to be published and has not been reviewed by fellow scientists - is highly significant because it suggests that all types of bacteria could become contaminated by genes used in genetically modified technology, including those that live inside the human digestive system.". Talk about hurried conclusions.

Isn't the phrase "genetically modified genes" kind of tautology?

I just want to point out that the Guardian article you refer was published in 2000. 12 years later, these putative results have not been published yet.

@MartaCz-C: I guess it was intended to be GMO's genes

No.
There is nothing special about a piece of GM DNA when compared to any other random piece of DNA. If this phenomenon happened at any detectable level, we'd have found eukaryotic DNA in bacterial genomes/plasmids long before the introduction of genetically modified crops. And that would be front page news in the field of horizontal gene transfer! Which has many talented people in it!

Do you have any references to back that up?

A basic molecular biology textbook plus one in microbiology would be sufficient to provide the details of the mechanisms described by Amy +1.

@Casebash You're asking me to prove a negative. I think the onus would be on you to provide evidence otherwise. For what it's worth, the wiki entry on horizontal gene transfer gives a pretty good overview of the mechanisms we know about: http://en.wikipedia.org/wiki/Horizontal_gene_transfer

Also, this textbook chapter: http://pathmicro.med.sc.edu/mayer/genetic%20ex.htm

Of far greater concern is the transfer of genetic material between bacteria in which antibiotic resistance is conferred. The firmicutes in your poop is not at all the same genetically speaking as the firmicutes in my poop. Polymorphisms and genetic transfer have changed members of the gut microbiome and that leads to differential extraction of energy from the ingested food and differential behavior with respect to antibiotics.

Apparently transfer between prokaryote and eukaryote (in both directions) has been detected:
http://www.biomedcentral.com/1471-2148/11/276/abstract
http://www.sciencedirect.com/science/article/pii/S0888754311001959
http://genome.cshlp.org/content/19/8/1441.abstract
Some numbers I'd like to know are the rate of pro<->eu HGT detection /versus the rate of looking for it/ (i.e. 3 confirmations in 300 attempts would be more impressive than 3 confirmations in 30,000 attempts).

Hi and welcome to biology SE. Please use the buttons above the editor window to insert links and it is better to include the title, authors and journal of your cited articles. That said, I have downvoted your answer because i) you link to 3 articles showing HTC between virii and eukaryotes (even if one of them seems to think that the Hz-1 virus is a prokaryote!) and ii) the existence of HTC between eu- and prokaryotes does not answer whether it would be possible for such a transfer to occur in the very hostile environment of a mammalian gut. In short, you're not answering the question :).


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ATP cost for gene expression
How would you estimate the number of ATPs required to transcribe, export and translate a single eukariotic protein?

Is this in bacterial or human? Transcribing a pre-mRNA and splicing are significant contributors to this energy cost calculation. Don't forget charging of tRNAs with amino acids.

Thanks, edited the question: I was thinking about eukariotes.

I'm assuming this wouldn't take into account the ATP cost of transcribing / translating / regulating any of the proteins required for transcription / export / translation?

@GWW, No: I assume the cell contains already all the required machinery.

The cost of transcribing and translating a hypothetical average gene in yeast has been calculated as 551 activated phosphate bonds ~P per second (Wagner, 2005).

The median length of a yeast RNA molecule is 1,474 nucleotides, and
  the median cost of precursor synthesis per nucleotide (derived from
  the base composition of yeast-coding regions) is 49.3 ∼P. With a
  median mRNA abundance of R = 1.2 mRNA molecules per cell and a median
  mRNA decay constant of dR = 5.6 × 10−4 s−1, the mRNA synthesis costs
  calculates as 49.3 × 1,474 × 1.2 × (5.6 × 10−4) = 48.8 ∼P per second
  and cell. This is a fraction 48.8/1.34 × 107 = 3.6 × 10−6 of the total
  RNA synthesis cost per second. The median length of a yeast protein is
  385 amino acids, with a combined biosynthesis and polymerization cost
  of 30.3 ∼P per amino acid. The median abundance is 2,460 protein
  molecules per cell. No currently available data allows a meaningful
  estimate of the median protein half-life, but a protein of an
  intermediate half-life (see below) of 10 h (decay constant dP = 1.92 ×
  10−5 s−1) yields an overall synthesis cost of 30.3 × 385 × 2,460 ×
  (1.92 × 10−5) = 551 ∼P s−1.

For your question about a single gene, the cost would be 49.3 * 1474 ~P for the mRNA and 30.3 * 385 ~P for the translation, which would result in around 84 thousand ~P. This is probably a very misleading statistic as you can transcribe multiple proteins from a single mRNA.
How the cost of mRNA synthesis and translation are calculated is described in detail in the paper. A large part of the cost comes from the synthesis of the basic building blocks, the nucleotides and the amino acids.

Wagner, A. Energy Constraints on the Evolution of Gene Expression. Mol Biol Evol 22, 1365-1374 (2005).


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Why do restriction enzymes tend to have an even number of bases in their recognition site?
When reading my textbook I noticed that in all examples but one from eight the recognition site was an even number of bases.
I wondered if this was just a co-incidence, so I took the data from this site for over a thousand known recognition sites and put it into a spreadsheet (XLS uploaded here).  The results are probably best summarised graphically:

Stats Test: Chi Squared Goodness of Fit.
Null Hypothesis:
There is no significant difference between the number of restriction sequences
that are an odd or even length.

|---------|----------|----------|-----|-------|----------|
|  Trait  | Observed | Expected | O-E |(O-E)^2|(O-E)^2 /E|
|---------|----------|----------|-----|-------|----------|
|  Odd    |    172   |  231.5   |-59.5|3540.25|  15.293  |
|  Even   |    291   |  231.5   | 59.5|3540.25|  15.293  |
|---------|----------|----------|-----|-------|----------|

Chi Squared Value = 30.586

P=0.05, 1 Degree of Freedom: Critical Value of 3.841

H0 rejected with 95% confidence (indeed with 99.9%+ confidence)

Can anyone explain or suggest why it is more common that restriction enzymes recognition sites have an even number of bases?
Updates

Expanded dataset to include new recognition sites from the resource that 96well linked to.
Removed all duplicate recognition sites leaving 465 distinct recognition sequences (my fault for not removing them in the first instance)
Ran stats test on the data
See Previous Version

I congratulate for your reflex: from the textbook to the database search. Nicely done! Your data is from a sample of some 1100 restriction enzymes. Rebase.neb.com reports some 6800 restriction enzymes. Are your conclusions still valid?

I think this is due to the over-representation of recognition sites with length 6:
data<-c(16, 16, 12, 12, 6, 6, 6, 6, 4, 16, 6, 6, 6, 6, 15, 15, 6, 6, 6, 6, 11, 11, 6, 6, 4, 4, 6, 6, 11, 12, 6, 6, 23, 23, 6, 6, 6, 6, 9, 12, 4, 4, 6, 6, 6, 6, 6, 6, 6, 6, 6, 7, 7, 10, 10, 6, 4, 6, 6, 11, 11, 9, 9, 6, 6, 6, 6, 5, 5, 8, 8, 6, 6, 8, 8, 6, 9, 10, 10, 6, 6, 6, 5, 5, 6, 4, 6, 6, 5, 5, 14, 14, 6, 6, 6, 16, 6, 6, 6, 6, 15, 15, 6, 6, 6, 6, 6, 18, 18, 7, 7, 11, 11, 20, 20, 6, 13, 4, 4, 6, 6, 6, 6, 6, 6, 6, 11, 6, 6, 7, 7, 6, 6, 6, 6, 6, 12, 6, 6, 10, 10, 23, 23, 7, 7, 23, 23, 12, 12, 6, 6, 6, 6, 10, 10, 6, 6, 8, 8, 6, 6, 35, 35, 11, 11, 7, 7, 6, 6, 9, 9, 8, 8, 16, 16, 6, 6, 17, 17, 6, 6, 6, 6, 23, 23, 6, 6, 4, 4, 21, 21, 12, 12, 20, 20, 6, 6, 6, 6, 6, 7, 7, 6, 6, 6, 6, 5, 5, 11, 11, 6, 11, 11, 6, 6, 5, 5, 7, 7, 11, 11, 11, 11, 6, 6, 12, 12, 6, 6, 6, 21, 21, 9, 9, 8, 8, 7, 7, 16, 16, 4, 4, 6, 6, 6, 6, 7, 7, 18, 18, 6, 6, 6, 6, 6, 23, 23, 7, 34, 34, 39, 39, 6, 6, 12, 12, 5, 5, 19, 19, 8, 8, 8, 8, 4, 6, 6, 6, 6, 6, 5, 5, 6, 6, 6, 6, 7, 7, 4, 4, 15, 15, 7, 7, 7, 7, 14, 14, 11, 11, 27, 27, 12, 12, 4, 4, 10, 10, 6, 6, 8, 8, 7, 7, 8, 8, 7, 7, 5, 5, 7, 7, 6, 7, 7, 6, 6, 8, 8, 39, 39, 6, 6, 12, 12, 8, 8, 7, 13, 13, 8, 8, 8, 8, 6, 6, 6, 6, 6, 6, 6, 6, 6, 6, 5, 5, 7, 17, 17, 17, 17, 11, 11, 15, 15, 6, 6, 6, 5, 5, 5, 6, 6, 5, 7, 7, 12, 6, 12, 6, 6, 6, 5, 6, 6, 7, 2, 5, 11, 5, 6, 4, 5, 7, 5, 4, 4, 6, 4, 5, 6, 7, 12, 6, 7, 7, 7, 6, 4, 4, 7, 5, 6, 6, 6, 7, 5, 12, 13, 5, 6, 6, 6, 5, 11, 11, 5, 6, 10, 5, 5, 11, 6, 5, 5, 6, 5, 6, 5, 6, 6, 7, 7, 6, 5, 5, 7, 6, 5, 6, 5, 6, 5, 5, 7, 6, 6, 6, 3, 5)
h<-hist(data, breaks=0.5:40.5)
df<-data.frame(counts=h$counts, mids=h$mids)
df$even <- (df$mids%%2 == 0)
ggplot(df, aes(x=mids, y=counts, fill=even))+geom_bar(stat="identity")

If you look at the histogram of lengths, there is no bias towards even recognition site lengths except for length 6.

I would guess that the followup question is why is this the case?

Interesting. Is there any motif in the length 6 sites?

I think you've found the natural bias that exists for recognition sites. The distribution is symmetric for 4-8 bases, and obviously cannot get much smaller than 4 without losing a great deal of specificity and as a consequence, not being very useful. At the other extreme, the larger the stretch of DNA required, the larger the enzyme must be which is expensive, and while you gain specificity you sacrifice cutting efficiency for things like methylation and secondary structure.

Surprising that there seems to be at least one restriction enzyme with a recognition sequence of length just 2!

Not sure why Larry Parnell was down-voted, he was not technically wrong.
Crystal structures of the most popular restriction enzymes are already known and can easily be found the Protein Data Bank or Wikipedia for graphical reference. Any stretch of double-stranded DNA makes a complete 360 rotation (about it's helical axis) in 10-10.5 base pairs. A 180 degree rotation would then be half of that ~5 bases.
A palindrome is not strictly necessary for a restriction enzyme site, but exploiting them has its advantages. Firstly, synthesizing one subunit of the enzyme requires half the occupied space of the genome, therefore translation is more efficient (2 products from half the instruction set).
Some restriction enzymes take advantage of rotational symmetry, such as EcoRI and EcoRII (both odd and even palindromic enzymes). They recognize the same GAA (in the case of EcoRI) or NNGG (in the case of EcoRII), coming from each of the two DNA strands. For notation purposes we write the recognition site of the positive strand only (since the DNA code is symmetrical). Asymmetrical recognition sites are explained by enzymes binding to DNA as heterodimers.
New England Biolabs has a very good review of the kinds of restriction enzymes and the variations in their behaviour.
http://www.neb.com/nebecomm/tech_reference/restriction_enzymes/overview.asp#.T0wqHYcgerY
To sum up, palindromes are often, but not necessarily, comprised of even numbers of bases (domains bind to 2n bases). Many enzyme recognition sites are palindromic in the literal sense (CATTAC) or biological sense (GAATTC) because there is an axis of symmetry that is used. Palindromic sequences are more commonly used because enzymes are composed of homodimers.

Could you expand on "domains bind to 2n bases" - I'm right in thinking that that is the reason for a preference of even bases?  Why is six so popular as mentioned in Biocs' answer?

If one half of a dimer recognizes "n" bases, and most of these enzymes (to the best of our knowledge of crystal structure/molecular weight analyses) are dimers, then that means they must be recognizing "2n" bases, which biases the total length of recognition site to be an even number.
From your very well collected dataset, there is an overwhelming amount of enzymes that recognize 5-7 bases, with the mode being 6. My inference based on crystal structures is that the recognition of ~3 bases is about a quarter turn of a double-stranded helix and is a conveniently recognized conformation.

Most restriction enzymes recognize palindromes, hence the recognition site of an even number of residues. Why palindromes? This allows for a vast increase in complexity (or rarity) from a DNA sequence standpoint, while requiring very little added complexity from the protein. In other words, if a protein domain recognizes 2 or 3 or 4 bp of DNA, simply adding a second copy of that domain to the peptide chain (via duplication of the DNA encoding that domain) does the trick. It is much easier to duplicate a module than to build a new or second (perhaps complimentary) module.

How does being even or odd affect a palindrome? What would be the probability difference between CATATG (6) and CATGATG (7) i.e. why are even palindromes more likely?

-1 for justifying your answer by defining palindromes as sequences with even number of bases. EcoRII recognizes the palindrome CCWGG (W being A or T).

@RoryM CAT TAC is not a palindrome. CAT ATG is a palindrome.

@GerganaVandova sorry I was thinking of palindrome in the common sense rather than the DNA sense, I'll make the edit!

The misuse of the word palindrome for these sequences is a real pet hate of mine - they're rotationally symmetrical not palindromic!

@RoryM: I have no idea why you're intepreting Larry's answer as suggesting that rotationally symmetrical - ;) - recognition sequences have to be even what he is saying is that the duplication of a single recognition module can produce a rotationally symmetrical recognition sequence and that this will lead to an even length recognition sequence.

Are odd numbered overhangs actually palindromes?  Somebody gave an example saying the following is a palindrome:
5'-CATGATG-3'
However, the opposite strand will have a C in the center position, that is, it will have the sequence:
5'-CATCATG-3'
In this example, the two center-most bases (G and C) are complimentary, but the two sequences are not strictly palindromic.  This should be the case whenever there is an odd number of bases in the overhang.
Another example, a 3-base overhang.
5'-GTC-3'
The complement will be:
5'-GAC-3'
Which is not a palindrome of the upper sequence.
This means that half of the time overhangs with odd number of bases cut by the same restriction enzyme from random fragments will not be complimentary (the 5'-GTC-3' overhangs can't anneal to other 5'-GTC-3' overhangs, only to the 5'-GAC-3' ones), and unless careful attention is paid to their sequences, odd-numbered overhangs in cloning projects can cause problems.
Even numbered overhang example:
5'-AATT-3'
In this case the opposite strand will have an identical overhang:
5'-AATT-3'
and will always have homology with other 5'-AATT-3' overhangs.
I suspect that the reason there are more even numbered restriction enzyme overhangs is to avoid this problem.

This is possibly better suited to be a follow-up question than an answer.

@MaximilinPress the last sentence does seem to address the problem, somewhat. To OP: I have added the 'add citation' banner, because the answer seems more like a long comment, or an educated guess at best. It's best to pen up science-based answers, including references to credible sources.

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


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What are the biochemical processes occurring when food spoils?
Let's assume for a minute that microbes themselves and their direct toxic products (i.e. endotoxins) aren't toxic to humans. Let's also discount any innate immune responses the body mounts against the invading microbe (i.e. inflammation and production of cytokines).
What happens to food molecules (mechanistically) as it spoils and what deleterious effects do these "spoil products" have on the body if ingested? I'm looking for compounds that can result from the spontaneous breakdown of food or the byproducts of microbial metabolism (that is NOT a "direct" toxin) that is harmful to the body.
For example, do the proteins in food break down into some toxic nitrogenous substance?

It is mostly oxydation as far as I know.

During putrefaction of animal tissue, lysine is decarboxylated into cadaverine and arginine is decarboxylated into putrescine. These compounds are deemed to be toxic.
A serving of meat contains 8 g of protein, corresponding to 640 mg lysine and a little bit less of arginine. Let's go straight and say that a spoiled meat serving contains 640 mg cadaverine and a little bit less of putrescine.
In rats, the acute oral toxicity for both polyamines is around 2000 mg/kg, let'assume that this is valid for humans also. According to these rough calculations, to have an acute toxic effect, a 70kg man that is resistant to the direct toxic effects of microbes, should eat 140 grams of cadaverine, corresponding to 218 smelly rotten meat servings.
[composition and toxicity data taken from wikipedia]

Cadaverine...hah. what a clever name.

Cadaverine and putrescine are legendarily foul-smelling. I do not think I could keep down 140 grams of pure cadaverine. That's like an orange, made entirely out of the solified scent of death.


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Density of neurons/cells in the mouse brain
Could someone point out some relevant papers or resources (an online DB maybe?) describing the density of neurons (or, more in general, of cells) in different areas of the mouse brain?

The newest and most accurate method (far more accurate than older extrapolating/manual counting methods (Stereology) and yielding some surprising results) to estimate number/density of neurons/cells in brains is Isotropic Fractionator to my knowledge. Using this keyword you find some recent papers, comparing different brain areas (cerebral cortex, cerebellum,...) among rodents:

Combining our estimates of total cell number and percentage of NeuN-containing nuclei in each brain region, we find that adult rat cortex contains ∼80 million cells, 40% of which (∼30 million) are neurons. In comparison, rat cerebellum contains more than twice as many cells (∼170 million), >80% of which are neurons (Table 1, top). Therefore, the adult rat brain contains almost five times as many neurons in the cerebellum (∼140 million) than in the cerebral cortex. When all of the brain regions are taken into consideration, the cerebellum thus accounts for more than one-half of the cells and ∼70% of all of the neurons of the entire rat brain (Table 1, bottom). Overall, we estimate that of all of the cells in the adult rat brain, 60%, or 200.13 ± 21.17 million, are neurons. Glial cells, therefore, contrary to common belief, are not the most numerous cell type in the rat brain.

Isotropic Fractionator: A Simple, Rapid Method for the Quantification of Total Cell and Neuron Numbers in the Brain
Cellular scaling rules for rodent brains
The Human Brain in Numbers: A Linearly Scaled-up Primate Brain

This article seems to be at least part of what you're looking for (although I can only access the abstract at the moment). They report that:

In this study the density of neurons and synapses was measured in three different areas (8, 6, and 17) of the neocortex of the mouse. [...] The total average was 9.2 × 104 neurons/mm3 and 7.2 × 108 synapses/mm3.

It would only make sense for the neuron and synapse density of the different areas of the mouse brain to be reported in the full-text article.

If someone can manage to obtain the information, can you report it? I'll edit the answer afterwards.


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Taxonomy for plasmids
I liked the question about F' and R plasmids.
What is a good resource that describes what is known about the functions and diversity of plasmids in the wild? Prokaryotes or eukaryotes?
Wikipedia points to five kinds of plasmids (fertility (F), resistance (R), col plasmids (offensive genes that let a bacteria kill other bacteria), degradative function plasmids, and virulence plasmids). There is a brief section on yeast/euk plasmids too. It seems like plasmids could actually convey nearly any gene. Restriction enzymes are found mostly on plasmids and they would seem to be defensive - maybe those are degradative.
Are there references that are more detailed and nuanced?

Not sure if generalized plasmid taxonomy is going to be relevant any longer. A lot of these old names were created before the exact sequence and function of the various DNA sequences were known. This is becoming especially true as synthetic biology allows us to mix and match parts of plasmids at will. If you want to dig through a lot of the old plasmid classification papers, a lot of them are published before 1980.
Back then I'd imagine conversations went more: "oh look, I found an R plasmid conferring ampicillin resistance", instead our more modern understanding: "oh, I have a plasmid conferring ampR via bla "
That being said, I think the more specific / specialized plasmid classifications will stick around for a while as they confer a lot of domain specific meaning.  Check out Ti and Ri plamids which have very specific meanings in plant pathogens and are used in the genetic engineering of them.  You'll notice that the classification for these plasmids comes from a lot more than a single gene.
There have been some modern attempts to classify plasmids, but rely a lot more on the structure / lineage of the plasmid rather than the payload genes (replication origin, size, etc.). Check out [1] for an example.
[1] Wang, et. al. 2009. Plasmid.

This makes a lot of sense - plasmid categories are probably highly artificial.  would you say that new categories of plasmid function are also being discovered as well as blurred boundaries between the plasmids functions already described?


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Determining potential protease sites within a recombinant protein
My expressed proteins are frequently truncated and I'm trying to figure out which bands are which. The first thing to come to mind is using PeptideCutter from ExPASy but there is just a data deluge of potential sites. I was curious what other strategies exists for determining the potential breaks asides using LC-MS.

Are you using E. coli for expression?  Using a protease-deficient strain (eg BL21DE3) is usually a good idea for testing this stuff.  If you still have truncations, then an analysis of the DNA sequence is in order.

I am using an E. coli expression strain but not BL21. Probably worth a try. I have tried using protease inhibitors and those haven't worked.

I should add that I've been using cell-free protein expression. However, the question at hand is completely independent of the expression system used.

LC-MS is certainly quantitative and will give you a definitive answer, but it is costly and requires access to such a machine.
I presume you're analyzing your protein based on western blotting.
The first thing you should always do is verify your DNA sequence is coding for the protein product you want. Once you're sure of this, the western blot will give you an indication as to where your protein is (approximately) being cleaved. Say you protein has a predicted size of 40 kDa and you see a band at 20 kDa, then your cleavage is somewhere in the middle of your sequence. There are a TON of proteases that could be potentially cleaving your peptide and you need to have a hypothesis as to what that could be. Your peptide could be cleaved by an endopeptidase (cutting within), or a carboxypeptidase (C-terminal cleavage) or an aminopeptidase (N-termainal cleavage). To go back to the 20/40 kDa example, it could be your peptide was cleaved N-terminally o C-terminally down to a size of 20 kDa, or that it was literally cleaved in the middle by an endopeptidase.
Something you may consider is the use of a general protease inhibitor cocktail (Roche makes a really good tablet product called Complete and Complete mini tab). These mixes have a bunch of general protease inhibitors which will stop most cleavage events. If you still see cleavage after using an inhibitor cocktail, you can reasonably expect that you are not inhibiting the protease with the tablet (thus narrowing down your search) and then you can better comb through Expasy's PepCutter data. You should also do a literature search to see what has been reported about your peptide or motif.

+1. Tried using the cOmplete tablets and still seeing problems. Do you have a good idea what's in the mix?

From the manufacturer: http://www.roche-applied-science.com/PROD_INF/BIOCHEMI/No1_99/pgs21-22.pdf
Notice that inhibition activity is not maximal, and decreases with time. Collection and manipulation of your samples must be done chilled and as quickly as possible to avoid degradation.

You could also try induction at lower temperatures to minimize chances of protein degradation. Cells such as Arctic cells (http://tinyurl.com/arcticecoli) are optimized for low temperature expression although BL21 works just as good for the proteins we express (upto 140kDa in size)

Good answer that is relevant to the problem but not relevant to the question.

I have tried lower temperature expression at 25C but still see issues. I could go lower to 10C.

@bobthejoe 10C is probably too low - try 16C first.

One way to learn more about how your protein is being processed/degraded is to add two different tags on either end of your protein. Express your protein, then blot with the two antibodies for your tags. The most likely scenario here is that one end or another is being cleaved and then degraded - you'll be able to tell which end by what tag epitope remains. And since you'll know the approximate size, you can narrow down the region where the processing occurs.
Small tags such as HA, cMyc, Flag or 6xHis work well because they are less likely to be cleaved off themselves.
The only way to know for certain how and where your protein is being truncated is empirically: either by N-terminal sequencing of the fragment(s) by mass spec or by mutating the putative processing sites and testing the expression again.

An epitope tag can tell you from a western blot where the protein is being cleaved (N-terminally, C-terminally, somewhere in the middle) without antibodies raised specifically for localized epitopes. Mutation of putative cleavage sites is a more specific way to go if you can identify what is cleaving the protein.


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What forms the human amniotic sac?
I am trying to understand the formation of embryonic disc for human and chick so the following question is related to here about formation of embryonic disc.
I had thought that the amniotic sac forms only from embryonic disc in humans, particularly of lateral mesoderm and ectoderm of embryonic disc. However, my thought seems to be wrong as my professor says that amnion form only for birds from embryonic disc, but from many different parts besides of embryonic disc for human.
What, besides the embryonic disc, forms the human amniotic sac?

In humans, the amnion (amniotic sac) persists from the primitive amniotic cavity1.  One side of this is formed from the cytoblast (a prismatic epithelium) and the plasmodioblast.  Together these two layers are the ectoplacenta or chorion.  They are also referred to as Rauber's layer.  These replace the lining epithelium of the uterus, whereupon internal cells undergo atrophy to create the amniotic sac.2
The other side of the amniotic sac is formed of the epiblast/ectoderm (internally) and the hypoblast (externally).  Within the epiblast the other layer is comprised of prismatic cells whereas the inner layer are flattened (the hypoblast/entoderm).  This double layer forms the bilaminar blastodermic membrane.2
I'd really reccomend that you borrow the referenced books from your university library as I found the diagrams much easier to understand than the text.

1 Gray, Henry. "Embryology: Formation of Membranes." Ed. Robert Howden. Anatomy Descriptive and Surgical. Ed. T. P. Pick. 15th ed. London: Chancellor, 1994. 90. Print. Colloquially Gray's Anatomy
2 Gray, Henry. "Embryology: The Ovum." Anatomy Descriptive and Surgical. Ed. T. P. Pick and Robert Howden. 15th ed. London: Chancellor, 1994. 82-83. Print. Colloquially Gray's Anatomy


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PDB Mining: Why Do I Find Atoms Less than 1 Angstrom Apart?
I am attempting to find potential Hydrogen bonds between Hydrogen donors and aromatic ring acceptors.  I do this by predicting the location of Hydrogens on residues and then calculating how far these Hydrogens are from aromatic rings.  If a certain Hydrogen is <7.0 Angstroms from a certain aromatic ring, then I take it under consideration: I form the N-H vector, which is the vector created by the Hydrogen under question and the Nitrogen in the backbone of the residue that the Hydrogen belongs to.  I test that this N-H vector is pointing toward the plane of the aromatic ring, and I also test that the point of intersection between the plane of the aromatic and the N-H vector is within 6 Angstroms of the center of the aromatic ring.
If all of these conditions are met, then I consider it a Hydrogen bond between the Hydrogen and the aromatic ring.  However, my data must be incorrect, because I am seeing situations where a Hydrogen is < 1.0 Angstrom from the plane of the aromatic.  Atoms should not be getting that close to each other.
I thoroughly tested my method by hand using an example situation where my code identified one of the sidechain Hydrogens on an ASN is 0.3 Angstroms from the plane of the aromatic of a TRP.  Unfortunately, I could not find any bugs.  You can find a PDF of this verification here.
Any suggestions on how my method might be flawed would be greatly appreciated.

Constructive criticism is far more useful than a down vote.

Try looking at the structure in question using a program like Chimera or PyMol, so that you can see how these programs interpret the direction in which the hydrogen should be pointing.

if you are using NMR structures you might be making the mistake of using several superimposed structures - it would be nice to develop with x-ray structures at less than 2.0 A resolution for starters.
Some of the models at low resolution can be sloppy, but submitted after 1994 or so will not have any center to center distances as that's when the x-ray structures used molecular models rather than just electron density rigorously and wierd steric violations in the structure should be rare.
Still, this might not be wrong - the hydrogen might be pointing right into the aromatic ring's center.  This has been observed quite a bit.  In such a case, the atom to ring plane distance might be very small.
You only need to worry if the center to center distance between atoms is less than the vanderWaals radius.   I would filter for atom center to center distance and see if you see violations.

Excellent idea.  Indeed, when I test for the distance from the Hydrogen to the center of the aromatic, I do not find any situations where the Hydrogen is within 1.0 Angstrom.  Indeed, situations where the Hydrogen is within 2.0 Angstroms is rare.  As for the NMR consideration, I did take care of that.  My parser successfully parses all of the NMR's MODEL's, and then it throws out all but the first one for the sake of this project

Now, I am left with only one problem: I expected to see most Hydrogen bonding situations between aromatic rings and sidechain Hydrogen donors.  To the contrary, I am seeing far more Hydrogen bonding situations between mainchain Hydrogens and the aromatic ring.  Can you think of any reason why?  I would expect the large backbone structure to be unable to get near to the small aromatic rings.

Glad you had already worked through a lot of this.  I would tend to think that main chain h-bonds will be selected for more strongly as the sidechains tend to change very easily.  Sidechains also tend to form stronger (i think stronger) aro-aro or pi-aro interactions and so there may be a tendency for important h-bonds to not from as readily.  Selection arguments are hand wavy, but maybe you can think of some clever stat test for this.


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What are the most important differences between HSP70 and HSP90?
Question originally asked on Quora. These proteins have many functional similarities, so why do cells need both to handle unfolded proteins?

For one, they have different molecular weights (HSP70 = 70kDa, HSP90 = 90kDa).

Often cells have multiple types of the same protein — this redundancy can have different effects for different requirements such as having proteins function under different physiological conditions, or providing specificity to a certain class of ligand proteins or so on.
But here, it seems like the two have some synergistic interaction, a tag team if you will.
 Wegele H, Müller L, Buchner J. 2004. Hsp70 and Hsp90 — a relay team for protein folding. Reviews of physiology, biochemistry and pharmacology 151: 1–44.
Unfortunately this article's full version can only be accessed if you're at a university or somewhere that has a subscription to some of the large research databases, but the abstract is free and it may provide more clarification.

Hsp70 and Hsp90 are not single proteins, but entire protein families. And those two protein families are not the only ones of their kind. There are different groups of heat shock proteins, Hsp70 and Hsp90 are molecular chaperones that assist in protein folding. There are five major families of molecular chaperones: Hsp100, Hsp90, Hsp70, Hsp60 and Hsps.
Hsp70 binds to nonnative, unstructured segments of proteins and helps folding them. Hsp90 doesn't bind to unfolded proteins, but to native-like proteins. Some substrates are also handed over to Hsp90 by Hsp70.
The whole area of chaperones and heat shock response is a very, very large and interesting field. I recommend to you to read the reviews below.

Vabulas, R. M., Raychaudhuri, S., Hayer-Hartl, M. & Hartl, F. U. Protein folding in the cytoplasm and the heat shock response. Cold Spring Harb Perspect Biol 2, (2010).
Richter, K., Haslbeck, M. & Buchner, J. The heat shock response: life on the verge of death. Mol. Cell 40, 253-266 (2010).


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Why does RNA adopt an A-form helix?
RNA is known to form an A-form helix, while DNA generally forms a B-form helix under physiological conditions.

From left to right: A-form DNA, B-form DNA, Z-form DNA. Image created by Richard Wheeler
The preference of RNA for the different conformation is supposed to be caused by the 2'-OH, my question is now how exactly this favors the A-helix? Which interactions are important and cause RNA to adopt an A-form helix?

There are many factors that produce effect in that interaction. You could have a first look in detail at http://www.sciencedirect.com/science/article/pii/S0022283605014439

The beta-form of DNA and the alpha-form of DNA are based on the pucker on the sugar ribose. As DNA doesn't have a 2'-OH, it can obtain both conformations.

RNA does not have this luxury. The steric clash of the 2-OH with the 3'-OH makes the B-form to be very unfavorable thus constraining the RNA to adopt the A-form. Incidentally this steric limitation is also why RNA structures are more stable (see https://biology.stackexchange.com/a/769/389)


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What can you tell about a person, having only their whole genome as information?
Well, the question is in the title, no explanation need.

I love the "only" part of the question :)

More every day.

Which level of certainty do you prefer? (If you want 100% certainty, you can tell less than if you were also considering risks and probabilities, and "Insecure knowledge", such as newest and speculative research findings. I hear that most things you could learn about from the genome are not that deterministic at all.)

I've had a little encounter with this question in the past few months so I'm updating here...
The overall answer is 'really a lot about some things, but not as much as you'd like to think about others.'    There is a scientific genome interpretation 'contest' that has been going on for the past few years called CAGI (Critical Assessment of Genome Interpretation).  This is meant to be a cutting edge set of challenges and its worth looking them over.  Last year there was in particular one challenge - answering questions about ten individuals given only a list of traits and their genome sequences.
It was not so easy it turns out - simply looking up variants and cross referencing them to the literature led to poor predictions.  Glaucoma, asthma, migrane, irritable bowel syndrome, color blindness, lupus, lactose intolerance are examples from a list of 40-odd contest questions.  If you register onto the site you can get some of the results or there is a paper reporting the results, only four labs tried the challenge and the accuracies were sometimes not great, topping off with Rachel Karchin's lab with an AUC of 90%.  Even some of the phenotypes you think are easy are not a simple lookup.  Genetics is not as pre-determined as we think.
While we are good at inferring our history and geneology on the other hand.  A reasonable example of this is the 23andme analysis.  They have a lot less information about you than a complete genome sequence, but they do seem to make the most of the 0.0001% of the genome that they do have.
They have a nice lookup of some of the hereditary disease data that is available.  "Increased risk of skin cancer" or for Alzheimers are only chances and its never clear how much your actual lifestyle has impact on 44 outcomes, but its there.
What they also have is your ancestral analysis which is crazy interesting. it shows where your maternal and paternal lineages come from and how strong.  they have a very pretty heatmap of the continents for this.  National Geographic's DB is probably more sophisticated and ancestry.com also does this.  This is pretty cool.
Also included things like curly hair, color of skin eyes and hair.
Like SimaPro says they are described in OMIM.  OMIM stands for 'online mendelian inheritance in man' and all the straightforward directly inheritable traits we know of are listed there.  its worth taking a look. There are not a lot of known single mutation genetically inheritable conditions.
If the genome sequence includes methylation, then some other traits which are epigenetic, not genetic but would also be determined from a genome sequence.  Most famous example of this is MEST which will give an indicator of whether you are a doting parent or not.

The first paragraph is (unintentionally?) misleading since you cannot extrapolate from the 0.00001% and conclude that the genome holds 10000-fold more information than 23andme is using: they are obviously picking their 0.00001% rather carefully. Given that there’s a SNP about every 100 bases, this then is the total increase of information you could gain about a single person – that is, a factor 100 over what 23andme is currently offering, not 10000.

@Konrad Rudolph: no, you would have a factor 100 if genes did not interact between each other, which is not the case. The effect of 2 SNPs at the same time may not be just the sum of those of the single SNP.

@nico True, as far as pieces of information are concerned. But as far as the information content goes, the numbers work out. Note that in information theory the information content is a logarithm of the number of different possible outcomes. Same here.

@Konrad Rudolph: sure, in terms of information content I agree with you.

Well - a lot if you knew what questions to ask...
The most obvious thing is sex: if you can find the x and/or y chromosomes.
Other things like skin color, eye color, etc, or hereditary diseases and all that would involve running the genome through some sort of database which had values for both pheno and genotypes so you could correlate some of those things and find those physical characteristics.
Scientists are discovering these correlations all the time.
You can go to http://www.ncbi.nlm.nih.gov/omim and search for something like eye color and find out more than you need probably.

The first place to look for answer to this is the 1000 Genomes Project. I feel that their efforts in data analysis offer lessons on what one can and cannot learn from such a dataset as posed in the question.
One obvious thing that is easy to overlook is a whole genome sequence will tell you the mid- to large-sized insertions and deletions in that genome. Any of those that are novel, private or extremely rare (take a word of your choosing) can be used to place that individual in a family tree (if other data are available) and to predict certain phenotypes (from the affected genes). This is also true for some SSRs - simple sequence repeats.
Good haplotype analysis from the genome sequence can predict ethnic ancestry - with a fair degree of accuracy. This is especially so as more genomes are sequenced and available for comparison. Haplotypes of the Y chromosome and mitochondrial genome, give paternal and maternal ancestry, respectively.
If the whole genome sequence gives accurate telomere lengths, then a prediction of age of the individual can be gained +/- 10 yrs.
Disease phenotypes or risk of disease is tricky. For monogenic diseases, this is more accurate and that accuracy falls off as the complexity of contribution to variance of the trait increases. At least a hundred loci contribute to risk of type 2 diabetes, but that risk is also age-dependent and so a young person may not have the disease but only be at increased risk. Furthermore, not everyone with elevated risk will acquire the disease and not all at reduced risk will remain disease-free - for that disease.
Lastly, if this analysis included epigenetic measures, such as methylation of the genomic DNA, then one can predict genome changes from environmental exposure such as tobacco smoking or air pollution. In addition, loci with differential methylation and showing a phenotypic consequence, such as elevated obesity status (BMI) or colorectal cancer risk, are beginning to be defined. This is in its infancy, but the data are growing.

Astonishingly, if they are male you can take a pretty good punt at working out their surname and narrowing down exactly who they are. There was a recent paper on doing this with anonymous genome data, discussed in a blog post here if you want a more accessible account.


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Is it possible to add a new sequence to a blast database without recreating it from scratch?
I have a new sequence and I want to add it to a pre-constructed BLAST database containing other sequences. The makeblastdb help does not give any hint on this regard.

You can use blastdbcmd to extract FASTA sequences from the BLAST database. Then you can append your extra sequence to the newly created FASTA file, and remake the BLAST DB using makeblastdb. Assuming you have a protein BLAST database in the current working directory, and a sequence you want to add to it in new.fa:
$ blastdbcmd -db proteindb -dbtype prot -out db.fa -entry all
    $ cat new.fa >> db.fa
$ makeblastdb -in db.fa -out newproteindb -dbtype prot

(Replace prot with nucl if you have a nucleotide database)

You may create a new database with the new sequences and create an alias linking the old and new databases with blastdb_aliastool


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Long-term-potentiation and memory. Where do we stand?
I was reading the answers to the question: How and where, in the human brain, are memories stored? and, as expected, LTP and LTD came out.
Every time I read about LTP/LTD there is always something that bugs me a lot.
When I read papers about LTP/LTD (OK, I should really say "when I browse", as I'm not exactly in that area of neurosciences, and that is probably the reason why I am confused by this) I always see these very neat experiments where stimulating a neuron in a certain way increases/decreases its further responses. Then, I look at the time axes on the graphs, or read the Methods, and I see that the LTP was induced and analysed few minutes after the stimulus.
So my questions are:

Is there clear evidence that LTP is involved in long-term memory (not counting 1 hour as long-term...)?
Has LTP/LTD been shown in vivo after long period of time (e.g. months).
For those who work in the field, is there a strong belief that LTP/LTD are the only phenomena underlying memory?

Is there clear evidence that LTP is involved in long-term memory (not
  counting 1 hour as long-term...)?  Has LTP/LTD been shown in vivo
  after long period of time (e.g. months).

This Journal of Neuroscience paper shows LTP in vivo measured out to one year:  Abraham WC, Logan B, Greenwood JM, Dragunow M. 2002. Induction and experience-dependent consolidation of stable long-term potentiation lasting months in the hippocampus. The Journal of neuroscience : the official journal of the Society for Neuroscience 22: 9626–34.

Thanks, that is what I was looking for. And bizarrely I even know the first author of that paper!

Good questions. I don't think that LTP has been (or will be) shown to be THE mechanism for long term memory. It is one of many mechanisms, all with different time courses, that contribute to the modification of synaptic efficiencies.
One mechanism not mentioned much anymore, but which I feel is absolutely crucial, is dendritic spine growth. Spines are malleable and formation/destruction of spines is probably very related to long term memory. But as always, the full picture is going to be a combination of mechanisms.
Here is a paper to have a look at:  Engert F, Bonhoeffer T. 1999. Dendritic spine changes associated with hippocampal long-term synaptic plasticity. Nature 399: 66–70.

Isn't this called engramming?

Really good questions. As the guy who brought up LTP/LTD in the question you referenced, I thought I would weigh in.
There is the traditional definition of LTP/LTD as an increased/decreased synaptic efficacy at a single synapse or in a single cell. As you've noted, this is unlikely to be the only phenomena underlying memory and sometimes it's hard to see how some of these mechanisms can result in memory on behavioral timescales.
Let me propose, therefore, that the term long-term plasticity is more relevant these days, as it can refer to a variety of mechanisms that relate to the ability of the nervous system to change in stable ways over time. Physiological mechanisms involving changes in protein expression include traditional LTP/LTD at single synapses, but also homeostatic plasticity and long-term changes in intrinsic excitability where the tendency of the cell to fire changes independent of changes in synaptic weighting. Some structural mechanisms include the growth of new synapses, new spines, and new neurons--synaptogenesis, spinogenesis, and neurogenesis.
In the end, it is all of these mechanisms (and probably more) at play. Note, for instance, that the plasticity may move through structural changes in the system. This means that the lifetime of LTP in one cell or at one synapse does not necessarily have to be the same as the lifetime of the memory itself. All that said, I think all plasticity mechanisms ultimately reduce down to a change in the ability of an input to elicit an action potential somewhere in the brain (known as EPSP-spike coupling). This is likely to be the basic underlying mechanism of memory.


text:
Is secondary follicle or Graafian follicle arrested in the second metaphase of oogenesis?
My note about which I am not sure:

Secondary oocyte is arrested in the metaphase II monthly after puberty until fertilization with sperm.

The following picture also satisfies my sentence: .
So you can see that the secondary oocyte that is arrested seems to be Graafian.
I know that Graafian develops during the second week.
I know that the secondary follicle is developing two days before ovulation.
It seems that the development of the secondary follicle ends and the formation of Graafian follicle occurs just before ovulation.
My lecture material suggests that Graafian follicle is arrested in metaphase II, since Graafian follicle is ovulated once in a month and waits sperm in tuba uterina infundibulum:

The answer to my question is not also clear from my previous thread:

An oocyte completes meiosis I as its follicle matures (during ovulation) resulting in a secondary oocyte and the FIRST polar body. After ovulation, each oocyte continues to metaphase of meiosis II.

Which follicle is arrested in the metaphase II of oogenesis?

Graafian follicle.
The secondary oocyte and thus Graafian follicle are arrested in the second metaphase of oogenesis.
After this the corpus luteum develops from the follicle during the luteal phase of the menstrual cycle.

Three follicular stages are recognised,
1) Primary/Preantral Follicles (with primary oocyte inside)
2) Secondary/Antral Follicles (with primary oocyte inside)
3) Pre Ovulatory Follicles (with secondary oocyte inside)
Pre ovulatory Follicles are formed ~36 hrs before ovulation at time of LH surge. This coincides with completion of Meiosis I and formation of the secondary oocyte. The secondary oocyte immediately enters Meosis II.
Appx. 3 hrs before ovulation, the Pre ovulatory Follicle (with the secondary oocyte inside) is arrested in the metaphase of Meiosis II and is extruded out of the ovary - a process called as Ovulation.
It comes to rest in the ampulla of the fallopian tube and here it is fertilised. As the sperm pronucleus is released into the cytoplasm, the Secondary Oocyte (now called the ovum) completes Meiosis II and releases the second polar body.
Hope this helps.

Hi Akshay. It would be great if you can cite a reference.


text:
Does chicken embryo form the disc called "discoblast" in the cleavage and blastulation of chicken?
This thread is related to my previous thread which is still unsolved mainly. I need to be able to compare chick and human cleavage and blastulation of zygote.
My friend says that

Chicken have discoblast, human not. That is the main thing to get the idea of the difference.

I know that chicken have yolk sac with yolk, while human have yolk sac without yolk.
My notes say that

Cleavage in birds is partial discoidal (embryo forms disc and cells called blastodisc are on top of the yolk).

So embryo forms some disc, lets call it #1. Some cells called blastodisc are on top of the yolk. So are the cells called blastodisc on top of the yolk and the disc #1?
What is the name of the given disc that chick embryo forms?
It seems that human is not forming the given disc. Apparently, the name of the given disc is discoblast, the thing he says is very important.

It is a mnemonic of two words: discoidal + blastula = discoblastula.
So it suggests that cleavage is partial discoidal in birds.
Embryo forms disc.
Cells called blastodisc are on top of the yolk.
It seems that the formed disc is blastodisc.

The cleavage in birds is restricted to the blastodisc lying on the top of the yolk towards the animal pole of the zygote, this type of cleavage is termed as discodial cleavage because the embryo forms disc and cells that are also on the top of the yolk.


text:
How to create a collection of anonymous sequences for teaching and testing?
I am looking for a large collection (>1000) of sequence files (eg. FASTA) from any real organism or a tool to create such a collection.
The sequence files would be used for teaching and for testing automation methods.
Students would be assigned one unique sequence file and asked to look at it (eg. using gORF) and to identify it (using BLASTn).
The sequence file would thus need to contain only the sequence data (no meta data about species or gene).
I would need an associated answer sheet.

I think you need to elaborate more on what you are trying to accomplish.  Furthermore, you may get more help on biostar

This question I asked in biostar seems related - Examples of DNA sequence motif sets for testing search algorithm

Here is the approach I ended up using, in part thanks to all the contributions here.
The associated R script is below or can be downloaded from:
BOLDS SEQUENCE RECOVERY
This creates 999 unique sequence files in plain text, with each sequence being identified to species level and few species being found across more than one sequence.
It also creates the matching answer key.
You can start at a random location to so that files change every year/group.
I used R to query the BOLDS database (Barcode of Life), to download a file and to split this huge file into separate sequences.
Here is the R script
rm(list=ls())

complete<-"http://services.boldsystems.org/eFetch.php?record_type=full&id_type=sampleid&ids=(*)&return_type=text"
write(complete, file="your location on disk")

rm(list=ls())

sequences.id<-data.frame("file.name", "recordID", "genus_name", "species_name")
write.table(x=sequences.id, file="sequences_id.csv", append=F, sep = ",", row.names=F, col.names=F)

set.seed(10)
start<-sample(1:1000, size=1)

i<-start
k<-1

while(k < 1000){

  sequences<-read.delim(file=complete, skip=i, nrows=1, header=F)
  sequence.compare<-read.csv(file="sequences_id.csv", skip=k-1, nrows=1, header=F)

  if(! is.na(sequences$V24)){
    if(as.character(sequences$V24)!=as.character(sequence.compare$V4)){
      writeLines(text=as.character(sequences$V55), con=paste(k, ".txt", sep=""))
      sequences.id<-c(k, sequences[,c("V3","V22", "V24")])
      write.table(x=sequences.id, file="sequences_id.csv", append=T, sep = ",", row.names=F, col.names=F)
      print("kept")
      k<-k+1
    }
  }
  i<-i+1
  print(paste(k,"/", i))
}

There are a couple key words in this question - anonymous and teaching. Yes, NCBI is a source for sequence data, but it is not anonymous (it is annotated, which means a student could also find it and copy/paste that annotation without doing the actual analysis). Notice, I am not assuming that the request is for human data. Now, if anonymous human data are needed, most available sequence data are anonymous but there remains that annotation issue: If it is annotated already, what will they learn?
A good alternate source for some human genome data would be from Complete Genomics. They have released anonymized (de-identified) data for at least 69 subjects. The question asks about 1000 sequences, but how large? This is an important consideration. Other details are also lacking in the question.
Another source may be the 1000 Genomes data, also human. If you're interested in plants, there are sequence data out there from ~98 different Arabidopsis thaliana accessions/cultivars/strains.

This is probably not the most elegant way but you could go to NCBI and search for nucleotide sequences from a given organism (eg., txid9606[Organism:exp] gives all sequences from Homo sapiens). Then you could use the Send to dropdown to download all the results as a compiled FASTA file.

NCBI also offers some APIs, which would make the process cleaner and allow better filtering.


text:
Which cryogenic vials and caps are ideal for storing glycerol stocks?
There are internally threaded, externally threaded, natural caps. For the vials, there are skirts, star-footed, and various bottoms. Why do any of these things matter?

I routinely store my stocks in plain vanilla 1.5ml eppendorfs and see no issues with it. I would be curious also to know what/why these special vials are "special"

Hence the question. Do companies just charge us more to make us think that they are "necessary"?

For any kind of frozen cell stock (for cell culture or bacteria) we routinely use freezing vials similar to these: http://www.sigmaaldrich.com/catalog/ProductDetail.do?D7=0&N5=SEARCH_CONCAT_PNO%7CBRAND_KEY&N4=V9255%7CSIGMA&N25=0&QS=ON&F=SPEC
These tubes are internally threaded with a rubber grommet to seal the lid and make the tube air and water tight. I suspect the air tight seal prevents condensation from entering the tube and freezing within the tube, but I've never had a problem with either choice of tube.
The tubes don't really matter at -80 C for a glycerol stab. Freezing cells in liquid nitrogen on the other hand has a huge impact when storing in liquid nitrogen. Some tubes are meant to be stored in the liquid versus the vapour phase of nitrogen and not matching the tube to it's intended storage condition can result in explosion of the tube.


text:
What is the closest species to humans in animal kingdom?
I presumed chimpanzees were the closest relatives of us. However, after watching this TED Talk, it seems bonobos are closer to us both in skeleton and behavioral similarity than chimpanzees. I once read in The Magic of Reality that we share quite a lot of common FOXP2 gene letters with chimpanzees, but there isn't any mention on similarity between us and bonobos in that book. Here is the excerpt:

You can tell that FoxP2 is the same gene in all mammals because the
  great majority of the code letters are the same, and that is true of
  the whole length of the gene, not just this stretch of 80 letters. Of
  the total of 2,076 letters in FoxP2, the chimpanzee has 9 letters
  different from ours, while the mouse has 139 letters different.   And
  that pattern holds for other genes too. That explains why chimpanzees
  are very like us, while mice are less so. Chimpanzees are our close
  cousins, mice are our more distant cousins. 'Distant cousins' means
  that the most recent ancestor we share with them lived a long time
  ago.
Monkeys are closer to us than mice but further from us than
  chimpanzees.   Baboons and rhesus macaques are both monkeys, close
  cousins of each other, and with almost identical FoxP2 genes. They are
  exactly as distant from chimps as they are from us; and the number of
  DNA letters in FoxP2 that separate baboons from chimps is almost
  exactly the same (24) as the number of letters that separate baboons
  from us (23).

So which species is closest to us, chimpanzees or bonobos?

Others have given great answers, so I'll just support their answers with this diagram and a little further background on how to think about relatedness in evolutionary context.

Since the author in this excerpt makes no mention of Bonobos, I would imagine that, by chimpanzee, he really meant the genus Pan, which includes both Bonobos and Common Chimps. But that does not necessarily matter.
Relatedness, in modern taxonomy, depends purely on common ancestry, not necessarily on similarity of specific traits. As others have explained, we make theories about ancestry relationships based on many criteria and logic, not just variation in one gene. In this tree, you should think of A, B, and C as theoretical, extinct common ancestors of the species below them in the tree. The list of apes at the bottom represent the relevant extant species species only. Bonobos and Common Chimps have a more recent common ancestor (C) than either Bonobos and Humans (B) or Common Chimps and Humans (B). This is why @mgkrebs said that both qualify equally as most closely related, because the common ancestor of each pair is the same.
Few would disagree that Bonobos do seem to be more similar to humans than Chimps are to humans, but that actually does not mean they are more closely related evolutionarily! Theoretically there are two ways this could happen even if Bonobos and Chimps are equally related to humans, either our common ancestor with Chimps and Bonobos also shared the traits we have in common with Bonobos and Chimps changed, or Bonobos and Humans happened to develop these similarities independently. The former is a simpler explanation, but either are possible for each similarity. Again, the key in understanding this is in thinking about the theoretical ancestor species, B and C. The only way Humans could be more closely related to Bonobos than chimps is if the common ancestor, B, was a descendant of C, which seems extremely unlikely and as far as I know is not supported by the genetic research in this area.
Our common ancestor with Gorillas (A) on the other hand is much "earlier" than our common ancestor with Chimps and Bonobos, so Gorillas are less closely related to use than the group containing Bonobos and Chimps.

Chimpanzees and bonobos are equally close to us in time (which is the opportunity for evolutionary divergence).  Our most recent common ancestor with either species is the same, as our line split from their common line about 6 million years ago (estimates are generally in the 4-8 mya range).
The split between the common chimp and the bonobo is much more recent, probably less than one million years ago.
So in terms of general closeness to us, these two species qualify equally.  For any particular gene, either one might be closer, as our version may be unchanged from our common ancestor, while one of the two Pan (chimp) species may be the same and one different.  For most genes that have differences, we can expect that we are the odd man out, while the two Pan species share a common gene.

Looking at one gene is not enough. Since each gene is free to evolve (or not), there can be different sister taxon relationships depending on the gene studied. A better estimate is made using several genes or combining the results of several analyses.
According to Purvis (1995), the extant sister taxon of Homo sapiens is a clade with the bonobo (Pan paniscus) and chimpanzee (Pan troglodytes), with gorillas as an outgroup.
So the answer is that neither is closer.
Purvis, A. 1995. A Composite Estimate of Primate Phylogeny. Philosophical Transactions: Biological Sciences 348: 405-421.

I'm don't have enough experience with this particular area to know whether experts have come to a consensus about this (although both your question and a brief literature search suggests they have not). What I do know is that when you begin looking at these types of questions at this level of detail, the answer that you get will depend on the characters used and the modeling strategy. The fact that you may arrive at a different conclusion when looking at a small number of morphological (skeletal) and behavioral characters versus genetic characters (sequence homology of a particular family of genes) is not surprising. Even when using the same characters, different methods (maximum parsimony vs maximum likelihood vs Bayesian) may give you different results.
So the answer is probably "we don't know for sure" (I'd be happy to be proven wrong here). But either way, the different conclusions you cite are not necessarily contradictory. Any conclusion will simply have to be justified in terms of the assumptions made, the data (characters) involved, and the method used.

please don't guess ans an answer.

As someone has mentioned in this question, the closest species not directly linked to the Homo genus are the chimpanzees and the bonobos. Around 6 million years ago, humans diverged from both chimps and bonobos, making them (as far as we know now) both humanity's last common ancestor. The last common ancestor directly connected to the homo genus, however, is Australopithecus for approximately 3-4 million years ago, as shown by the Little Foot skeletal system in South Africa .


text:
Displaying nucleotide at a single position from RNA-seq reads in a BAM file
How do I display a single nucleotide position from reads in a BAM file? I have been looking at variation using samtools mpileup, but I want to actually just display the nucleotide at the position I am interested in. This seems like you should be able to do it but I can't figure out how.
To be clear I have a BAM file of a bunch of reads. I'm looking to do something like samtools magic reads.bam chr3:10000 where I get back something like:
T T T T T T T T T T T T T T T T T T T A A A A A A A A A A A A A.

I just want to sanity check the output of bcftools by actually looking at the base calls.

Your best option is to create a tab delimited position list file:
chr3    1000

You can then use the samtools mpileup -l pos.txt option
-l FILE      list of positions (chr pos) or regions (BED)

This file can also be in bed format if you are interested in specific regions


text:
How long can E. coli stocks be stored at -20°C?
I'm volunteering for a biohacker lab - biocurious in Sunnyvale.  The have a pretty good set of equipment - gel boxes, incubators, but they don't have a -80°C freezer yet.
I'd like to set up some glycerol stab stocks of some E. coli strains, but i'm told even untransformed (i.e. no plasmid) bacteria will get funny after a while at -20°C.  It would also be good to hear how long you can keep a plasmid transformed strain at -20°C.
Can anyone be specific about what happens and how long it takes when you store them at -20°C for long periods of time?
I have done a little reading for competent cells and I guess they stay active for about 4 days without a -80°C freezer.

so i'm getting that everyone thinks that the bacteria will simply die at -20C.  I asked this question because I was wondering if it was apocryphal.  Everyone 'knows' that -80C is the best temperature for glycerol stabs, but i'm wondering if its 100% necessary, esp over something like 3-4 months.  Some good suggestions though.  I will try it with some stabs and get some protease deficient strains as well.

You can also start keeping transformed cells (with some drug resistance marker) at -20°C, and plate some of them every day and see how quickly they will lose the plasmid. You can do the same with untransformed competent cells.

some great points here.  If its okay i'm going test the answers to see who is right before picking one.  if anyone is in the peninsula area of the sf bay area, you are welcome to come to biocurious and check it out. ;)

I might do that:) Can you email me (gvandova@stanford.edu) with some more details about it?

looks like it is 1 year at -20C..http://www.opsdiagnostics.com/notes/ranpri/aguidetobacteriapreservation.htm

Mother nature's got you covered. -136 ˚F (-93.2 ˚C). Locate you samples: in a small valley between two peaks on the East Antarctic Plateau. ~Dec 10 2013; ;) Maybe not what you were looking for, but for this one point in time you could have just put your stabs out on the back porch. Now that is a cold day! http://blogs.scientificamerican.com/observations/2013/12/10/found-the-coldest-place-on-earth/

Hey @GerganaVandova - i was just looking this over and realized I never did contact you. Shortly after this question my second child was born and I did not get back to biocurious much after that :/  Apologies!

Gergana covered the "why" part of your question. +1
If all you have at the moment is a -20°C and mostly what you want to store is E. coli harboring plasmids, I'd recommend preparing plasmid DNA and storing that at -20°C. The DNA will stay stable in the medium term, and you can re-transform into E. coli once you've got your -80°C freezer up and running. And ideally you want both DNA stocks and bacterial stocks on hand - you never know when a freezer will go down.
For your untransformed strains, you could make agar stabs or plates to store at 4°C. I would re-streak them every 1-2 weeks, grow overnight at 37°C, and then put them back in the fridge.

I think that cells get damaged at -20°C not because they are stored for a long period of time, but because they undergo cycles of thawing and freezing (the ice crystals that form damage them).
I never keep cells at -20°C. I store them at -80°C in 50% glycerol. The idea behind the glycerol is that it serves as cryoprotecor. It forms hydrogen bonds with the water molecules and thus it disrupts the water-water hydrogen bonds. As a result, it disrupts the crystal formation of ice, and the cells don't freeze. The freezing temperature of glycerol is −37.8 °C (60-70% glycerol in water), which means that at -20°C glycerol does not freeze and thus it cannot protect the cells from damaging from ice crystals formation.

A good -20°C freezer should not cycle between freeze-thaw. This is a problem with frost-free freezers, warming up to near O°C. Indeed, the glycerol is the trick for preservation of membrane integrity. Decades ago, researchers stored their bacterial cultures at room temperature, in the dark, in what were called stabs.

When I said a freeze-thaw cycle, I meant when you take the frozen sample from the freezer, do something with it (it thaws), and then put it back in the freezer (it freezes).

This is an interesting answer - if it's only the ice crystals, then one could use 40% glycerol and the solution would freeze solid while still being mostly just as good a cryoprotectant. http://en.wikipedia.org/wiki/Glycerol_(data_page)#Freezing_point_of_aqueous_solutions I tend to think that there may be some more specific answers out there though...  I'll try a 40% glycerol stock in LB (or whatever it takes to freeze solid and see how this works.

@shigeta I didn't know that proteases are active at -20°C (leonardo's answer). But it is worth the shot (at least short-term) if you don't store the cells too long.

The answers already posted about glycerol being a cryoprotectant and ice crystals are correct. The other thing to consider is that enzymes still retain some activity at -20°C and though the reaction rate is slower, the cells will degrade until they are no longer viable. I always store glycerol stocks of bacteria at -80°C.
Many common E. Coli strains commercially used for cloning are not protease-free and these are the enzymes that are responsible for cellular degradation.

Which enzymes are active at -20degrees?

Proteases. Just edited my post.


text:
What is the molecular mechanism of cystine bond formation?
My bioinorganic and redox chemistry isn't up to par and I'm having a very difficult time understanding the chemical mechanism behind cystine bond formation. Most reactive environments are actually reductive whereas an oxidative environment is important for cystine bond formation. How does the oxidative environment oxidize the thiols, rearrange the orbitals and form a disulfide bond?

Orbital rearrangement, etc, seems to be a bit of stretch for this site.

@DanielStandage: why? Biochemistry is definitely on topic

@bobthejoe: why do you think oxydation of the thiols is peculiar? In other words what would be so "special" about the "environment", and of which environment are we talking about?

I hope that biochemistry is on topic. Most reactive environments are actually reductive. An oxidative environment (via a periplasm or a chaperone) is important for cystine bond formation. I'm just curious about how those conditions are regulated. I hope that is clear.

@bobthejoe: OK, now I get your point. I don't have a precise answer, but I would like to hear one!

@bobthejoe You're asking a completely different question in your comment, please make it clear in your question what exactly you're looking for. Are you interested in how this happens in cells? Then I'd drop the part about rearrangement of orbitals as it's only confusing the question.

If your question is with respect to a eukaryotic cell, the di-sulfide bridge/bond is formed in the rough endoplasmic reticulum which is an oxidative environment (unlike most other organelles which are reductive). This paper may be of relevance to you:
Pathways for protein disulphide bond formation - Frand et al, Trends Cell Biol. 2000 May;10(5):203-10.


text:
PCR amplification and error propagation
Say I have one dsDNA that undergoes normal PCR (where amplification is exponential).  If there is a mistake, say a G is swapped for an A during the 2nd round of replication, what percentage of the final DNA will have the mistake if there are 12 more rounds?
I was talking to my teacher about this today.  I say 33%, my teacher said 12.5%.

I will format it to .ppt as soon as I have more time! If something is not readable, please let me know!
I have to point out several things:

The fraction of incorrect DNA molecules does not depend on the number of cycles (as long as the number of cycles is higher than 2).
There is one exception: if the mutation occurs outside of the region to be amplified (for example at the 4th molecule, downstream of the region of interest), the total number of wrong DNA molecules will be 0.

Your teacher is indeed correct.
In the first round you would get two identical molecules of the dsDNA.
In the second round you would get 3 identical molecules and one molecular with an A substituted for a G in one of the strands.  ie.
No error (3 of the 4 molecules):
------G-------
------C-------

One mismatch (1 of the 4 molecules):
------A-------
------C-------

So there are a total of 8 strands of DNA after the second round and one of those strands has the mismatch.  1 / 8 = 0.125 = 12.5%
In round 3 you would have 8 dsDNA molecules and only one of those 8 dsDNA molecules would have the mismatch.


text:
Can any protein be phosphorylated?
I am working with an Arabidopsis mutant with an F-box protein knocked out. It has been shown that F-box proteins targets must first be phosphorylated (Skowrya et al., 1997). I have heard of phosphorylation sites, but I can't find out whether every protein has them. Can any protein be phosphorylated?

Skowyra, D., Craig, K.L., Tyers, M., Elledge, S.J. & Harper, J.W. (1997) F-Box Proteins Are Receptors that Recruit Phosphorylated Substrates to the SCF Ubiquitin-Ligase Complex. Cell. 91 (2), 209–219.

Phosphorylation can occur on specific amino acids only, what you have called phosporylation sites. These amino acids are Ser, Tyr, Asp, Thr and His. In theory any of these amino acids may be phosphorylated, but in reality it may not actually occur for a number of reasons. Some of these are because of the change in overall charge of the protein which can influence the 3D conformation, or the amino acids are not accessible to specific kinases, etc. If you ask for the purposes of doing a Western blot, then the antibody specification sheet should indicate whether a phosphorylated form exists and there should be a reference to the literature describing this modification.

One important thing is missing in the other answers: not only phosphorylation will happen only at selected aminoacids, but it will not happen at all of those.
So, not all of the Ser/Thr/Tyr of a protein can be phosphorylated because they could be structurally unaccessible to protein kinase and because they need to be in a specific motif in order to be phosphorylated.
The Human Protein Reference Database, for instance, lists the phosphorylation motifs for many Tyr and Ser/Thr kinases.

I actually said this in my response. ;)

@leonardo: you did not speak about phosphorylation motifs which are a requirements for phosphorylation. You don't need just a Ser, you need a Ser with specific aminoacids around it.

Ah my mistake. I had touched on conformation being an issue but not the motifs. Thanks for the correction.

@leonardo: no problem, I re-added the conformation problem just for completeness :)

Phosphorylation requires exposed serine, threonine, tyrosine, or histidine residues (in eukaryotes).  This is because the transfer of phosphate groups to proteins is mediated by a class of proteins called kinases.  Kinases can have broad or specific activity.
This review ought to have most of the answers to your questions :
http://www.cell.com/abstract/S0092-8674(06)01274-8

I like Nico's response best +1. I did find an interesting review on phosphoarginine and phospholysine - the list of possible phosphorylation sites grows. Not only is it the spatial context of the amino acid to be (or not) phosphorylated, as Nico writes and Leonardo implies, but also the temporal. Are the target protein and protein kinase co-expressed? If you're looking at specific tissues (root, flower, leaf) and the kinase in question is only produced in seedlings, maybe that potential phosphorylation won't really occur.

For one of the most comprehensive databases of protein post-translational modification (including phosphorylation, methylation, acetylation, ubiquitination, etc.), check out PhosphoSite. You can find links to sequences, diseases, motifs, publications, antibodies, mass spec experiments, structures, you name it.