text:
Which range are the Therapeutic Indices of Currently-Prescribed Benzodiazepines in?
A compound's therapeutic index is defined as:
$$TI = \frac{LD_{50}}{ED_{50}}$$
What range are the therapeutic indices of benzodiazepines in? For the method of administration I would prefer intravenous as with it one can estimate the therapeutic indices via other routes. Of course for humans it is difficult to find LD50's but a rat/mice LD50 will suffice provided the ED50 is for the same respective species. Such benzodiazepines I am interested in are (Although I will accept your answer if you just give me an estimate of the therapeutic indices of the drugs in bold, the others are just icing on the cake):

Alprazolam
Brotizolam
Clonazepam
Clorazepate
Diazepam
Estazolam
Flurazepam
Lorazepam
Lormetazepam
Midazolam
Nitrazepam
Temazepam

Wikipedia quoted this article as a reference for its statement about alprazolam having a therapeutic index of 100:1, although I can't confirm that, since I can only read what's immediately available in that link. There's a figure saying less than 100:1 for diazepam though.

Thanks, an answer is better than none at all. I'm afraid I probably won't accept your answer. I will, however, upvote it, since I know just how tricky it is to find an answer to this question, hence why I'm asking it.

A first pass answer is 662-4342.
The Therapeutic index is calculated from rat tolerances; LD50 is a rare number to see for human beings(!).  The toxic dose of a compound can vary widely over a class of compounds as well as does the Effective Dose (ED) midpoint (ED50).
But I found this book reference:

Benzodiazepines have a wide margin of safety based on experimental studies in animals.  In the rat for example, the lethal dose of alprazolam is 331-2171 mg/kg. The effective therapeutic dose for alprazolam is 0.5 mg/kg which results in a therapeutic ratio of ... 662-4342.

The page goes on to say, at rather great length, that human evidence is that despite the fact that this is a popular class of drugs to try to kill yourself with, there are relatively few human deaths, which is pretty ultimate a judge for toxicity, though you could look for FDA warnings on the class of drugs.
I would bet that you can find relatively toxic benzodiazepines, but for the ones that pass screening they seem pretty benign as a class.  There are other issues with these drugs rather than mere toxicity - they are quite addictive, so therapeutic index is not an ultimate judge of the adverse effects of a drug.
BTW taken literally, this is a hard question you ask.  If you are expecting a compiled list of all such drugs, cross referenced to their Therapeutic index, that is really more like the work for a paper rather than 25 points of stackexchange karma IMHO.  On the other hand maybe someone has this information at their fingertips...

Is the effective dose of 0.5mg/kg for humans or rats? In order for a therapeutic index to be a fair estimate both the lethal and effective doses have to be for the one species.

That was for rats as well. The therapeutical index is, in general, not suitable for quantitative comparison though. The figures may even vary wildly for one substance. It's at best a rough indicator of safety margin. Have you looked into an alternate definition, TD50/ED50? Defined for humans, where TD50 is the dose that produces a toxic response in half the test subjects.

Yeah I know of the therapeutic window but the problem is that we know more about therapeutic indices for other drugs so it's a more useful comparison for me.

That information would have to be dug up through pharmacology texts or the literature.  Maybe someone could know that off hand and post, but you need to name specific drugs... there could be dozens of drugs in the class.  That's asking a lot.


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What are the most well-understood vocal animal languages?
Just as the Bee dance, for a "language", I mean that there are vocal pattens, or "grammar".
In the nature, there are many intelligent animals like human beings.
Bird songs, whale songs, dogs?
In fact, bird vocalization includes both bird calls and bird songs. There are Parrots, hummingbirds and songbirds,...
I wish to know the most well-understood vocal animal languages. Which species?

"most well-understood" is rather subjective, this question doesn't really fit the Q&A format well. It is also cross-posted on CogSci, so I'm closing it here on this site.

@MadScientist Why subjective? Apis mellifera is one of the most well studied animals, if not THE most. Is this subjective? I know of some African monkeys that we know have distinct words for "snake", "eagle", "hierarchy above", "hierarchy below". I know some whales can repeat exactly the same song one and a half hour long. I think the question is very interesting, and it will have different answers here than it will on CogSci, and probably very interesting answers. So I vote to reopen it.

source: Stack Exchange

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Studying changes in DNA for causes of cancer
First of all let me say that I'm not into Biology myself... but I have a question for those of you who are.
From what I've read, cancer is caused by 'faulty' DNA that behaves abnormally. Mutations can occur for various reasons, ranging from simple 'copying errors' to environmental factors.
I was just wondering whether current research has dealt with storing DNA of people or animals (e.g. rodents) and observing how it changed, especially for cases where it eventually resulted in cancer. Something along the lines of this, but I'd like to know what research has actually been done so far.

I tried to comment but what I wrote is too long, so here it is as an answer of sorts.
If I understand the question, you are asking: has anyone done a prospective study where they store the DNA of individuals and then later, when some of these individuals get cancer, have a look for mutations that are associated with that cancer. In fact this is done all of the time, but it isn't necessary to store any DNA because the cancer patients will have the normal sequence in all of their non cancerous tissues - only the cancer cells will have the relevant mutations. See here for a recent example of this type of study that made the news.
Your BBC link is to a story in which the genomes of people will be analysed to determine if there are genetic factors which predispose them to cancer (and other diseases), but these won't be the mutations that actually create the cancerous cells.

There is a tremendous amount of information relating to mutations of genes and how they produce cancer.  In fact, this is multi-billion area of research.
What is even more interesting, is the aneuploidy hypothesis of cancer, in which it is chromosomal abnormalities, rather than mutations in single genes, that lead to cancer.
I suggest reading about it, there was an article by Peter Deusberg in a 2007 Scientific American article.

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Confusion related to gene expression
I have a set of gene expression data downloaded from http://www.ncbi.nlm.nih.gov/geo. I have two sets of data, one is the raw probe intensity data set in the form of CEL files. Another is processed txt file.
Here is a link for example
http://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE7904
I don't know what kind of processing is done to generate the processed data from the raw CEL files. For example in the above link, I have processed data like
SOFT formatted family file(s)
MINiML formatted family file(s)
Series Matrix File(s)
I don't know what sort of processing is done to generate those files. Any suggestions? I can read the raw CEL files in matlab but what sort of preprocessing am I supposed to do?
I also want to map these probe its to gene ids and get the corresponding gene. How can I accomplish this? I know matlab. But I am bit confused about the terminology and all. Any suggestions?

this website converts affy id's to various gene id codes if this is the sort of thing you are after

@rg255. I didn't get how the processed data is generated. As you can see here http://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE7904 there are two data sets. One is raw CEL files and the other is Series Matrix File(s). How is this processed data generated. Also if I look at the number of rows. In the case of Cel files, the number of probes is huge 1354896. But when I look at the series matrix data it is around 50000. I didn't get it. Any suggestions?

@rg255. Also I didn't get the link you said this website. I didn't find the link attached

oops, http://david.abcc.ncifcrf.gov/conversion.jsp - as for the rest of your question I wouldn't like to answer as I am not an expert

If you click through to some of the samples in the study, eg. GSM194397, GSM194398, etc. it mentions under the "Data processing" section that "The data were analyzed with dchip with default normalization settings".
You can learn more about dChip over at the dChip website.

Thanks I will try that. However it looks like I will need some CDF files as well. Where can I get them? In particular I need the CDF files for chip type HG-U133_Plus_2 and HG-U133A. Any suggestions? I looked a couple of places but looks like I have to pay for it. It was supposed to be free I guess

I suggest you use the limma package in bioconductor to analyze the data. I'm not sure why one study has data from two chips -- it's unfortunate. You'd first have to study each batch separately, then do some meta-analysis to combine the results from each together. If you're not comfortable programming, you might try Gene Pattern, though I've never used it. Lastly: you should likely post bioinformatics related questions over on biostar.

I need to get the gene id for the corresponding probe it. For that I need the CDF file. So I was actually wondering where I can get them?

There are other ways to convert id's to genes. I'd be surprised if you can't query BioMart to get that answer (look in the ID Converter tab). It also looks like the DAVID website has a conversion tool that may get you what you need.

The methods used for the processign of the date should be mentioned in the Methods section of the paper the data come from (here it is Richardson AL, Wang ZC, De Nicolo A, Lu X et al. X chromosomal abnormalities in basal-like human breast cancer. Cancer Cell 2006 Feb;9(2):121-32. PMID: 16473279). I quote from the paper:
"RNA extraction, cRNA synthesis, and hybridization to Affymetrix Human Genome U133 Plus 2.0 Arrays were performed as described previously (Signoretti et al., 2002 and Wang et al., 2004). Raw expression data obtained using Affymetrix GENECHIP software was normalized and analyzed using DNA-Chip Analyzer (dChip) custom software (W.H. Wong and C. Li, http://www.dChip.org/). Array probe data were normalized to the mean expression level of each probe across a sample set. Where indicated, tumors were classified as BLC or non-BLC on the basis of their expression array characteristics, using dChip hierarchical clustering analysis as previously described (Matros et al., 2005 and Wang et al., 2004). Comparisons between results obtained on BLC or BRCA1 tumors, non-BLC tumors, and normal breast samples were performed using the dChip “Compare Sample” function. A threshold of 1.2-fold overexpression in BLC and BRCA1 tumors was applied with 90% confidence. Of 1271 gene probes that map to the X chromosome, 60 satisfied these overexpression criteria with a range of fold difference from 1.35 to 5.11. The false discovery rate (number) of 1000 permutations was as follows: median, 0% (0); 90th percentile, 3.3% (2). Of the 60 probes, 19 were redundant (two or more probes mapping to the same gene) and excluded, leaving 41 gene-specific probes for use in the expression plot of Figure 5B. The complete gene expression array data set is available on the NCBI GEO database (accession no. GSE3744)."
Since this is a standard Affymetrix human genome chip, it's CDF should either be embedded in most packages used to analyse this kind of data (limma is an industry gold standard example ;-), or downloaded from the Afymetrix website. Please note that the CDF itself may be quite old and it's annotation not always accurate any more (I have no experience with human data), so it's worth using an updated CDF (see Dai et al 2005).


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Why are pili more common in Gram negative bacteria than in Gram positive?
Although pili have been observed in some species of Gram positive bacteria, the preliminary research that I have done indicates that pili are significantly more common in Gram negative bacteria.
Is this an accurate assessment of the current body of scientific knowledge? If so, do we know (or have any leading theories on) why this is the case?

Pili were at least discovered more recently in gram-positive bacteria.
Pili are formed differently in gram-positive and gram-negative bacteria.  Here's a pretty good review on the differences: http://www.ncbi.nlm.nih.gov/pubmed/18953686
Researchers seem to think that pili might be involved in adhesion/formation of biofilms (adhesion is important for pathogenicity).  Certain types of pili are specifically involved in gene transfer. This is a good review on pili and pathogencity in gram-positive bacteria: http://www.nature.com/nrmicro/journal/v4/n7/pdf/nrmicro1443.pdf. If you can't access it, let me know, and I'll do my best to summarize.
EDIT:
The Wikipedia entry for pilin interprets the second paper I linked as saying that pili are more common in gram-negative bacteria and that pili are implicated in pathogenicity.
Straight from the paper:
Prevalence:

Over the past five decades, several distinct pilus types
  have been identified, most of which were described
  and characterized in Gram-negative bacteria.

Function:

A common feature of Gram-negative pili, however, is their role in adhesion to eukaryotic cells. It has been
  proposed that bacteria use these structures to form an
  initial association with host cells, which can then be
  followed by a more ‘intimate’ attachment that brings
  the bacterium into proximity to the host-cell surface.

and

Pilus-like structures on the surface of Gram-positive
  bacteria were first detected in Corynebacterium renale,
  by electron microscopy. More recently, surface
  appendages were reported to be present in Actinomyces naeslundii
   and were subsequently found in other
  species, including Corynebacterium diphtheriae, Streptococcus parasanguis (Streptococcus parasanguinis), Streptococcus salivarius
   and Streptococcus sanguis (Streptococcus sanguinis). Finally, in the past
  year, pili were also characterized in all three of the
  principal streptococcal pathogens that cause invasive
  disease in humans — group A Streptococcus (GAS; that
  is, Streptococcus pyogenes), group B Streptococcus (GBS;
  that is, Streptococcus agalactiae) and Streptococcus
  pneumoniae — in which they have been shown to
  have key roles in the adhesion and invasion process and
  in pathogenesis. [emphasis added by me]

this doesn't precisely answer the question.  are they more common in gram negative bacteria, and if so, why?

@shigeta: see my edited answer.  Given the diversity of pili structure and function (see the first article I posted), I don't know if you'll get a "why" that explains pili prevalence in gram-negative bacteria.

of course perfect answers aren't always going to happen with biology, but it helps the site to give the answer here.  thanks!


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What's the therapeutic index of Pethidine (Meperidine in the US)?
What's the therapeutic index $\left(\frac{LD_{50}}{ED_{50}}\right)$ of Pethidine (Meperidine in the US) via the intravenous route for humans or whatever species you can find? The therapeutic window $\left(\frac{TD_{50}}{ED_{50}}\right)$ for humans will suffice if accompanied by the therapeutic index for pethidine via IV of another species.
Pethidine is an opioid painkiller with approximately $\frac{1}{7}-\frac{1}{8}$ the strength of morphine and a definite influence on monoaminergic neurons.
I imagine it'd be in the neighbourhood of 30-100, like most opioids that have been implemented in medicine.

I don't understand the downvotes. This seems like an answerable question.

Tell me about it... It would be nice if those that think I've done a poor job in asking this question could tell me what I did wrong.

I have not downvoted but perhaps people feel this is a medical question not a biological one. A biologist is unlikely to know the answer, a pharmacist or a surprisingly knowledgeable doctor would.

Pharmacology is a biologica/biochemical pursuit, not just the purview of medicine or pharmacy.

If so, where am I supposed to ask this question as far as stackexchange sites? This biology stackexchange is best suited for me to ask this question.

In any case I created a new tag to suite this question.

Just go ahead and ask. Remember that anyone can answer on SE, so someone with the answer may come along. Feel free to publicize your questions in other venues. Unfortunately, I'm not sure what those venues are for pharmacology questions.

Maybe someone was reading your question upside down and thought they were up-voting....

Sounds about right. I've been trying to get a pharmacy stackexchange site started but that's going to take at least a few months even if it gets a relatively large amount of support.

Closest I reached:

Source: Page 105, The Complete Recovery, By Anthea Hatfield Google Book


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Does the NMDA antagonist, Memantine that's used to prevent Excitotoxicity cause brain damage?
Does the NMDA antagonist, Memantine that's used to prevent excitotoxicity in Alzheimer's disease (AD) and Dementia with Lewy Bodies (DLB) cause brain damage? I know that in rodents NMDA antagonists like ketamine and phencyclidine (PCP) cause a form of brain damage known as Olney's lesions. Despite Olney's lesions never being directly observed in primates like humans, permanent brain damage from chronic ketamine exposure has been observed in adolescent cynomolgus monkeys.1
Reference List

Sun, Lin; Qi Li, Qing Li, Yuzhe Zhang, Dexiang Liu, Hong Jiang, Fang Pan, David T. Yew (2012). "Chronic ketamine exposure induces permanent impairment of brain functions in adolescent cynomolgus monkeys". Addiction Biology. doi:10.1111/adb.12004. PMID 23145560.

Yes, it does cause lesions in rats, although I don't think it's been observed to do so in humans.
See: http://www.druglib.com/druginfo/namenda/description_pharmacology/

Thanks, mate. That's helpful for me, of course in human studies would be more helpful but c'mon there's no such studies on the most widely-used NMDA antagonists like ketamine and PCP so I can hardly expect it for memantine -- a drug I had trouble finding evidence even in rodents.

@BrentonHorne Apparently, memantine might cause hallucinations in humans (N=3): http://jnnp.bmj.com/content/78/5/546.1.short

What do you mean by N=3?

N is commonly used to refer to sample size (at least among the biologists that I've spoken with).  The article I linked is a case study of 3 individuals.  My point is that this is an extremely small number of individuals with documented adverse affects to memantine. If you want to read the article, you can read the 1 page preview (as an image)...the whole article happens to be 1 page.

Ah, thanks, I assume you're talking about hallucinations because it makes the link between its NMDA antagonist activity and potential brain damage more clear by demonstrating that it has one of the effects characateristic of NMDA antagonists, i.e. hallucinations.

Yes, I did mean the hallucinations.


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What is a Mitochondrial Eve and Y-chromosomal Adam
I've read this question:
What kind of event would cause the current Mitochondrial Eve to be replaced by a new one?
And the values in Wikipedia about Mitochondrial Eve and Y-chromosomal Adam and I am not sure I understand the concept behind them.
When talking about a maternal unbroken linage and a paternal unbroken linage, is the meaning that Mitochondrial Eve had a linage of only women reaching every person today, and the same with Y-chromosomal Adam and men? So that other humans that lived in their age, didn't have an unbroken line of linage consisting of only same sex descendants?
How is the specific "identity" or time period of those persons is known, assuming that researchers didn't check the Mitochondrial DNA or Y-chromosome of every single living human being today, how can they know for sure that there aren't some tribes in the Amazons, which have a direct female linage to Mitochondrial Eve's sister?

I'd note that if they have a direct female lineage to Mitochondrial Eve's sister then that would make their mother the true Mitochondrial Eve. Whilst the estimate may be inexact, there must be a mitochondrial eve at some point in our ancestry.

When talking about a maternal unbroken linage and a paternal unbroken linage, is the meaning that Mitochondrial Eve had a linage of only women reaching every person today, and the same with Y-chromosomal Adam and men?

Kind of, yes. The concept behind Mitochondrial Eve and Y-Chromosomal Adam has to due with cell replication and basic genetics.
The Mitochondria ("Power Generators" of the cell) are directly inherited from mother to daughter cell. That is, new cells do not start with 0 and are forced to make them de novo. Some Mitochondria are replicated before division and some are 'given' to the daughter cell to start with.
Because of the unique way that Mitochondria are inherited, when it comes to creating new humans it means that only the Mother's Mitochondria are given to any children she gives birth to regardless of the sex of the child. So I, being male, have my Mother's Mitochondria - which the same as my Grandmother's - and the same as my Great-Grandmother's - etc. etc. etc. until you can trace that line back to "Mitochondrial Evel" whose Mitochondria are the origin for all humans on the planet and are inherited Maternally.
In that sense it is an unbroken lineage of genetic information as inherited from mothers, though not exclusive to women.
"Y-Chromosomal Adam" is a similar concept - Women do not have a "Y"-Chromosome, so every Y-Chromosome men have today had to originate from somewhere. It's trickier to do than tracing Mitochondrial DNA because while Mitochondria are inherited exclusively from mother to child, there could have been multiple Fathers to Mitochondrial Eve's children and other similar complications that muck things up.

How is the specific "identity" or time period of those persons is known, assuming that researchers didn't check the Mitochondrial DNA or Y-chromosome of every single living human being today?

For "Y-Chromosome Adam" it's still more or less in the data-gathering phase as far as I was last aware. It's not as concrete as "Mitochondrial Eve", which has a very solid body of work supporting her.
The "identity" is known because the more in common a person's genes have with the genes of another, the more closely they are related. The researchers took a lot of genes from very divergent populations, and compared them. The researchers were then able to tell which portions were common among all of the Mitochondrial DNA, which had changed the most, and which had changed the least from each other. Using that data, you can construct a rough image of what the originator (or LCA - Last Common Ancestor) had for their own genetic makeup.
You can construe a time period because portions of your DNA (either Mitochondrial or Nuclear) acquire/tolerate mutations at an expected rate - say 1 mutation every 1.5 Million Years. So as long as you have a rough idea of what your LCA's genes looked like (see above), then a modern day sample with 5 mutations is 7.5 M.Y. away from the LCA.

how can they know for sure that there aren't some tribes in the Amazons, which have a direct female linage to Mitochondrial Eve's sister?

They didn't when they started testing. That's part of the fun of the project.
The "Out of Africa Hypothesis" rose in prominence partially due to the findings of the research. It basically posits that the ancestors of humans evolved once in Africa, and then spread from Africa to the rest of the world. Because the Mitochondrial Eve research concludes that "Eve" was around before we spread throughout the rest of the world, it implies that every ancestor already had Eve's legacy when they started migrating into Europe, Asia, the Philippines, etc. Thus every human on Earth, no matter where they are today, shares traits with Mitochondrial Eve.
For the Amazonian tribes, they would have inherited it from their ancestors in North America, which were ancestors of tribes in Asia who crossed an ice bridge into North America, and the tribes in Asia came out of Africa.

Just to note that you don't always get the mitochondria from the mother, sometimes you get some from the father too (but this is very rare.) This doesn't really change anything other than the "unbroken line of mothers" aspect.


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Is there an "Adam and Eve" for each new trait during (not just human) evolution?
Do new beneficial mutation evolve simultaneously and independently with several individuals of the same specie or only with one specimen who then transfers it to all of the specie?
In other words, when we talks about common ancestor we usually mean a specie, so for example, all earth dwelling reptiles, birds and mammals are the descendants of Tiktaalik (to the best of my knowledge), and we know that Tiktaalik was the first (or one of the first) species to come to land, but was there a first individual Tiktaalik who went to the land, like Tim Minchin's Tony the fish.

Imagine being the first dude ever to have feet...
Imagine you're a fish, and you are swimming in the ocean ... and suddenly out of nowhere Tony just goes "I'm getting out, I'm freezing, I'm just gonna pop out for a bit, maybe lie on my towel, I'll bring you an ice cream"
-"Don't bother ,Tony, you are a freak! Tony is a freak, let's go, let's ostracize him for being different"

It seems to me that from the same species of ancestors there are several descending distinct species, so it seems to me that that that there was a species of reptiles that at some point some individual of that group developed Dinosaur features and traits and whose descendants became Dinosaurs and then Birds, and another individual of that specie or a descending specie (not necessarily at the same time) developed mammary glands whose descendants became mammals, and other individuals who didn't mate with those two whose descendants remained reptiles.
Was there ever a first dinosaur with feathers who is the father or mother of all birds, or the first fish with lungs, the first monkey with opposable thumbs from whom all apes descend, or the first mammal that decided to go for a swim and all dolphins, whale and seals are his direct descendant of that first swimming mammal?
And also for traits that don't lead to speciation, was there a first human who was lactose tolerant, and all the lactose tolerant people on the world are his/her descendants, or the first person with no wisdom teeth, the first person with slanted eyes, and the first person with red/blond hair?
*- If there are gross inaccuracies and/or over-simplification in my understanding of the evolutionary process, this is because I'm not a biologist, and I would accept happily any correction.

I think I'm with Michael Kuhn. The concepts between the questions are very close. Ultimately the answer is 'Yes' - there's an "Adam and Eve" or "Parent" to every extant species today, but it's probably not like you'd expect. They were simply able to pass on their trait more successfully than others.

I'm not talking about speciation here, like the question you reference to, but about new mutation/features. I'm aware that the first dinosaur with feathers is still a dinosaur, and the first fish with lungs is still a fish, the question is, was there only 1 first fish with lungs, who's the ancestor of all creatures with lungs, or were there several fish that all developed lungs independently, and then interbred between themselves to produce all lunged creatures, but there is no 1 specific ancestors to all, but several different individual ancestors.

@IlyaMelamed - So you're talking about different ways traits evolve. Convergent, divergent, etc. If you can find a way to make that clear in your question, then I'd take away my Duplicate vote since there doesn't seem to be a good question covering evolution of traits.

@MCM, I hope it's clearer now

If you a new mutation which is recessive, although there is one progenitor, the phenotype might not be realised until many several generations later when the first homozygous offspring is born. In this case you don't really have an Adam and Eve. You could even have the first exhibitions of the phenotype occur simultaneously in different populations.

@IlyaMelamed by the way, my understanding is that many dinosaurs had feathers, including modern birds.

For most traits, there is no clear line where the trait is present or not. Exceptions are traits that occur with a single mutation, like lactose tolerance. So to pick one example, you had a population of fish that had a lung-like organ, and that by genetic drift that organ got more lung-like in a sub-population. So for most traits, you have a combination of mutations that lead to the traits, and these mutations need not occur in a direct line of ancestors. (I.e., a descendant will combine independent mutations from its parents.)
So there is no "Adam and Eve" in the sense that these are the only ancestors of all living individuals of a species. This would be a terrible genetic bottleneck! See this illustration, which comes with the estimate that the population size of the chimp-human ancestor was around 100,000.

You ask for "a first dinosaur with feathers who is the father or mother of all birds". It's possible that feathers arose multiple times from the same feather-like structures, or only once. But it is clear that this first dinosaur is only one of many dinosaurs that lived at the same time, and who are also ancestors of all birds.
Further reading: Lactose tolerance evolved several times independently (see "Convergent adaptation of human lactase persistence in Africa and Europe.").


text:
Why does throat 'burn' when you've got a common cold?
Apart from pain when swallowing, there's also a persistent feeling of throat 'burning' in the same area.
What causes it and why does it feel that way?

Could you describe the burning sensation some more? Is it located the same spot as the soreness?

@Brian well yes, pretty much the same.

If you've been sneezing or coughing a lot it is probably micro abrasions in the throat. I don't recognize any other burning sensation.

My common cold started with sore throat, with no running nose or coughing.

I guess it is because the throat is sore, thus probably inflammated. Inflammations are hot. Though, I am not a doctor.


text:
Finding proteins in DNA sequence
I have to do a task for a university task and I need to understand some things
before figuring out how to do it.
The task is the following:

Find matches of known proteins (DNA-PolyI,II,III) to the specific E.Coli DNA, sequence.

I downloaded in FASTA format the protein sequence of DNA-Poly3 DNA-Poly1 of E.coli (strain K-12) and the entire DNA sequence
of the E.Coli.
I've studied a bit on-line and using the BioRuby gem and the Ruby programming language I wrote a program that translates DNA to protein sequence.
Then I tried to match the known DNA-Poly3 sequence but it did not match. After searching a bit on-line again, I learned about ORF and
and the 6 possible reading ways of each frame. The longer, in terms of codons, ORF conformation is chosen but there's no way of telling for sure that the protein was made using
this frame.
Then I've read about TATA boxes, but I can't use those since they can be found only in Eukaryotic and Archaea.
So how should I proceed in order to solve this problem: How can I prove that the DNA-Poly3 gets produced by a specific area (gene) in the DNA sequence?
Thanks for your time,
ps. Insights and hints are very much welcomed as this is just the tip of the iceberg for me and I'm very willing to study bioinformatics :-)
EDIT: This is an update for info requested in relevant answer
The files I have used are the following:
➜  Bioinfo  ruby dogma.rb
----------------
DNA Length: 4639675
gi|48994873|gb|U00096.2| Escherichia coli str. K-12 substr. MG1655, complete genome
----------------
DNA Poly-1 sample: 928
gi|16131704|ref|NP_418300.1| fused DNA polymerase I 5'->3' polymerase/3'->5' exonuclease/5'->3' exonuclease [Escherichia coli str. K-12 substr. MG1655]

You can download them here: E.Coli DNA and E.Coli DNA-Poly1.
NOTE: My sample protein is DNA Polymerase I (and not 3).

Do you need to write a program that does this or will existing tools be OK?

I should write the program and put up a web interface.

OK, but do you need a totaly new algorithm or can you use existing tools like BLAST? If this is a computer science course, you should implement the algorithms yourself (I guess), if it is a bioinformatics course you should not reinvent the wheel. There are many, very, very good, programs that do what you need already. Is it OK to integrate them into your web interface?

Yes I should not (by any means) re-invent anything. I can use algorithms already in use. I know there are very sophisticated tools for the job. What I really want to know to begin with is how it works, then I can use ready algorithms.

To give a clear example, all my data comes from ncbi (BLAST). How come that I can not match it? I've tried out all 6 possible conformations of ORFs and still I can't "produce" a sequence that matches the DNA-Poly3! :-/

Aren't the RBS and Shine Delgarno sequences a much better search term?

Hello @bobthejoe, could you give me links and offer a quick hint about why are they better suited?

The DNA sequence you have is the entire genome of the bacterium. It contains a multitude of ORFs coding for all sorts of proteins. What you need to do is use BLAST, or much better, exonerate as described in my answer. Translating the entire genome and then trying to find a single protein is both incredibly inefficient and prone to errors as well as very hard.

I am trying to create some results. After reading the manuals - it has an incredible amount of options - I used the following cli: exonerate -Q protein -T dna -E -m protein2genome:bestfit dna-polyI.e-coli.fasta e-coli-K12.fasta  and I'm waiting for it to finish. My hardware is a macbook air i5 1.7 Ghz with SSD.

You are only interested in the best result here since that will be the coding gene. You do NOT want exhaustive (E), that is only useful for highly divergent sequences, not when working on the same species. Try this: exonerate -m p2g dna-polyI.e-coli.fasta e-coli-K12.fasta it took about 2seconds on my laptop and the first result is what you want.

IMPORTANT EDIT : In your particular case, if you are working with bacterial genes, splicing is not an issue since bacteria do not have introns. I am leaving the information here since it may be useful to someone else. However, I recommend you focus on the UTRs since they are probably what is causing you problems.

There are three things that could be causing you problems. I will briefly touch on each one. I will talk about all genes, bear in mind that bacteria have no introns so any discussion of splicing and/or introns and exons is not directly relevant to your problem.
1. UTRs
Untranslated Regions (UTRs) are sequences at the beginning and end of a gene that are not translated into protein. UTRs are regions that are part of the original genomic sequence, they are also part of the mature mRNA (indeed, UTRs are sometimes modified by splicing events, they are exons not introns) but they do not get translated into protein. To illustrate, have a look at this simplified representation of an mRNA molecule:

Only the green exons will make it into the final protein. Introns are spliced out and UTRs are not translated.
Therefore, if you translate the entire gene, you will not get the correct protein.
2. Reading frames
Genes are read in words of three letters (the codons). The sequence ATGTGTACCTGA has six possible reading frames (three on each strand) which can be read and translated as follows:

5'3' Frame 1
    ATG TGT ACC TGA
     M   C   T  Stop

5'3' Frame 2
    a TGT GTA CCT ga
       C   V   P

5'3' Frame 3
    at GTG TAC CTG a
        V   Y   L

3'5' Frame 1
TCA GGT ACA CAT
 S   G   T   H

3'5' Frame 2
t CAG GTA CAC at
   Q   V   H

3'5' Frame 3
tc AGG TAC ACA t
    R   Y   T

DNA is double stranded. The sequence of one strand is complementary to that of the other, therefore if you have one strand you can infer the sequence of its complementary one. Genes can be found on either strand, the two are equivalent biologically. However, sequencing projects choose one of the two strands (randomly) and call it the plus (+) strand and then save all sequences with respect to that strand. This means that sometimes the genomic sequence that you download from a database might be the complement of the actual sequence you are looking for.
3. Names
I once heard someone say in a conference that

Biologists would rather share a toothbrush than a gene name.

While that might be a little exaggerated, naming conventions vary between research communities and species and databases. So, are you sure that you have downloaded the correct gene? Where did you get it from? How did you identify it? Does the sequence also contain up/downstream regulatory regions, promoters, enhancers and the like? If you post the exact sequence you are attempting to use I can give you more specific help.
For example, the first 20 hits when searching for the E. coli DNA Polymerase 3 in ncbi's nucleotide database, are whole genome shotgun sequences. These do not correspond to the gene sequence you are looking for. They are huge pieces of the genome (or even the entire genome) that will contain your gene and many others. Look at the Tools section below for suggestions on extracting your gene from the whole genome.

4. Splicing (irrelevant to bacteria)
Another possible problem is splicing. Lets start with the basics, the process of producing a eukaryotic (bacteria have no introns) protein from a genomic sequence is summarized in the image below (modified slightly from here):

Transcription begins at the transcription start site (TSS) but not all the transcribed sequence is translated into protein. First, the introns are spliced out of the mRNA to produce the mature mRNA (other things like capping and poly-A addition also occur but are not relevant here). So,the mature mRNA contains the exons of the coding gene. This means that a linear translation of the gene's sequence will not correspond to the protein produced. You will need to take splicing into account.
Also, bear in mind that splicing will change the reading frame.
Now, if the sequence ATGT were spliced at, for example, AT/gt (most splice events cut/join at GT/AG sites) and joined with the sequence agATTATT, the resulting (spliced) sequence would be (the splicing process will remove the gt from the first sequence and the ag from the second):
ATATTATT

As you can see, the reading frame has now changed. Where before, in the first reading frame, we had the codon ATG, the canonical translation initiation codon, we now have ATA which codes for isoleucine (I). I hope that is clear,  the main point is that splicing can change the reading frame.

5. Tools
OK, that was the background. Now, what you will need to do is use existing programs that model splice sites and can correctly align a protein sequence to genomic DNA. My personal favorites are exonerate and genewise. On a Debian-based Linux distribution, you can install them with this command:
sudo apt-get install exonerate wise

Then, to align the protein to its gene do:
exonerate -m protein2genome -n 1 prot.fa dna.fa > out.txt

or
genewise -pep -pretty -gff -cdna prot.fa dna.fa > out.txt

In my experience exonerate is (much) faster but genewise is a little more accurate. I usually use exonerate if I am dealing with a whole genome and genewise if I only have a few kilobases of sequence. Both are very good and both will be able to align a protein to its genome of origin.
I will not explain all these options because that is beyond the scope of this site. Have a look at their documentation (which is quite good and clear) and if you still have problems, you could ask a question on our sister site, Bioinformatics Stackexchange
Alternatively, you could link your web application to the ucsc genome browser BLAT service. Click here to see the results when aligning the human
DNA-directed RNA polymerase II subunit RPB1 protein.

Thanks for the answer! I will study the answer in a couple of hours when I'll have time! Thanks for your time!

Just to let you know, the way to give thanks on the Stack Exchange network sites is to upvote the answer and, if (and only if) it answers your question to accept it. Thanks are always appreciated of course, I'm just letting you know how these sites work.  Anyway, you're very welcome :).

The OP asked about an E. coli protein and the E. coli genome. I don't think his problem is anything to do with splicing.

@AlanBoyd ah, you make a good point. Probably down to the UTRs then. I'll edit my answer.

Hello, thanks for the explanation it was enlightening, I will try work on it tonight. Do you know which is the main procedure by which the protein is aligned to the DNA? :-) Thanks again!

Please re-read my answer, I made a substantial edit. You can ignore everything I said about splicing. As @AlanBoyd very correctly pointed out, if you are working on bacteria it is not a factor. As for aligning proteins to DNA, use exonerate or genewise as described. You can also use a genome browser like the ucsc or ensembl.

Sure, already did! Thanks again for the tips!

@terdon I modified my initial question and added some more specific questions! Thanks for your help, it's very very much! You saved me quite some time!

@terdon However, sequencing projects choose one of the two strands (randomly) and call it the plus (+) strand and then save all sequences with respect to that strand. This means that sometimes the genomic sequence that you download from a database might be the complement of the actual sequence you are looking for.What are the serious implications of this method?Please elaborate.Thnx

@Raghavakrishna what do you mean? What method? If you mean the two tools I mentioned, exonerate searches both strands by default and genewise does so if you give it the -trev flag.

@terdon I meant the methods used by the sequencing projects to  choose one of the two strands (randomly) and call it the plus (+) strand and then save all sequences with respect to that strand

@Raghavakrishna I still don't understand what you're asking. Sequencing projects sequence one strand and call that the + strand and then extrapolate the sequence of the - strand. If you have a question about this, please ask a new question so you can explain it clearly and get a full answer.

For what its worth - I have replicated what you are trying to do using a Python script. This is not elegant, but I just wanted to check for you that it is possible, and that there really is a match.
pseudocode is

take the genome sequence
make a reverse complement sequence
for each of the two DNA sequences, for each of three reading frames:
translate the DNA into a single string of amino acids with "*" at stop codons
split the string at "*" characters, call these words
find the first Met residue in each word, the string from that Met to the end of the word is an ORF
if the ORF is >99 (arbitrary cut off) put it in a big list of ORFs
now have a list of all ORFs in all 6 reading frames
search this list for a match to the polI sequence (I actually just looked for the first line in the fasta sequence).

The hit is identical to the entire polI sequence in a CLUSTAL alignment.
Note that this algorithm does not detect any ORFs that cross the breakpoint in the linear sequence representing the circular genome of E coli. Also assumes all initiator codons are ATG/Met but I seem to recall some E.coli initiation codons are GTG/Val

Rather than doing it all from scratch, if you had your own instance of BLAST, you would make a blastable database of your e.coli sequence, and do tblastn, with your putative polymerase protein sequence as the query.
This would find the best matching sequence in the genome, and will work even if there are a fair number of differences between the protein you gave it, and what your DNA sequence actually translates to.


text:
Can the immune system stop plasmodium from being active?
Suppose a female Anopheles infected with Plasmodium bites someone and transmits Plasmodium to their body.
Can that person's immune system be strong enough that it can kill the Plasmodium before it starts entering into red blood cells to rupture them?

There appear to be some references here: http://www.bio.davidson.edu/people/sosarafova/Assets/Bio307/ruturakhia/page03.html

The short answer is yes, at least with P. falciparum. Even if they're not taking anti-malarials, not everyone who gets bitten by an infected mosquito becomes symptomatic (I don't know about RBC entry, that may happen too quickly). Unfortunately I don't have the references handy, which is why this isn't a formal answer, but this information came from a professor studying the immune evasion techniques of the parasite.

@MattDMo hmm!. I always tell my friends who curse the medical science to not to eat anti malarial tablets when they get malaria. I tell them that one infected with malaria will 100% die if they don't eat antimalarial tablets which is a medical science creation. so MattDmo am I right when I say that to support medical science to my friends who it. I mean if malarial patient doesn't take antimalaria, will he die?

@MattDMo why don't you answer it instead of adding comments. I want to upvote you for this answer that I can't unless you answer it

The experience of a former adviser of mine was that some people who are not taking anti-malarial medication can still resist infection with Plasmodium falciparum, one of the parasites responsible for malaria. The parasite's life cycle includes stages inside the Anopheles sp. mosquito, and they are transmitted to human host via the female mosquito's saliva when she bites to have a blood meal:

(from Wikimedia)
There is currently a great deal of research going on to understand the biology of the parasite and how it typically evades the immune system so well. Proteins expressed by the parasite can undergo antigenic variation, meaning that different forms are expressed at different stages of the life cycle, and just as the immune system may be forming a response to one version, it is down-regulated and another takes its place. This is one reason why making a vaccine has proved to be so difficult, as unlike the influenza virus, for example, a single strain of Plasmodium can alter its immunogenic appearance and escape antigen-specific immune responses.
While malaria is responsible for a high degree of morbidity - sickness - it is not necessarily fatal except in some cases, frequently when the victims are young, already ill, or are otherwise immune-compromised, or when the infection moves to the central nervous system and the brain. I can't speak from personal experience (fortunately!), but I have been told by numerous friends that malaria is something you want to avoid at all costs. The sickness can last for weeks in some cases, and apparently is quite debilitating at its worst. Brain fever can be deadly, and in many third-world countries and among the poor the lack of good medical care can lead to complications. There is the additional factor that the family has to care for the sick, and if the ill person is a wage-earner that can mean the difference between food on the table and going hungry.
So, your friends are not going to die (probably) if they don't take their tablets, but there is no reason not to take them unless you have strong side-effects, and in that case the doctor may be able to find you another one. Other common-sense measures include always sleeping with a bed net, putting screens in windows, covering exposed skin during active mosquito hours, wearing repellant if available, and eliminating places where standing water can collect, stagnate, and provide a breeding area for mosquitoes.

This paper estimates the mortality rate of untreated malaria as 1.8% (http://www.ncbi.nlm.nih.gov/pmc/articles/PMC3021416/).  The lifetime cost in terms of protracted illness are not considered.  There's a good reason that antimalarials are useful and worth taking in the future. If you haven't taken them before, that's another thing.


text:
Resuspending Cells from a filter plate
My experiment involves pulling down cells on to a filter plate for a variety of assays. However, I want to ensure that my cells are alive so I am trying to resuspend them so that I can do an accurate cell count. However, I am unable to close off the mass-balance which suggests that the cells are either getting through the membrane or getting stuck.
Any suggestions on the appropriate way to fully resuspend cells? I'm currently using a P200 to mix the solution.

Have you tried trypsinizing them? Depending on whether they're suspension or adherent, and the time they've been on the membrane, you might be able to detach them just like you would from a culture dish/flask. Also, are you using a vacuum to adhere them to the membrane, or just allowing them to settle? Too much pressure may be getting them caught in the pores, although theoretically the pores should be much too small for them to fit through...

@Mattdmo, these are suspension cells so trypsinization will kill them. I'm using a vacuum manifold. I'm fearing the same thing which is why I'm doing this experiment to see if I can complete th mass balance

Resuspending is hard due to attachment/entrapment in the filter and liquid retention around the edges.  Consider using optical live/dead staining on the membrane?   If you are concerned about pass-though you can also try and capture the fluid that comes through the membrane and give it a hard spin in a centrifuge.  Look for a tiny cell pellet on the outside bottom (mark the caps).  Or a simple and cheap dye for viability or counting that would be great on a filter


text:
Does GTP-γS (GTP gamma S) bind all GTP-binding proteins?
I've just read an article Rab10 GTPase regulates ER dynamics and morphology - Nature Cell Biology 15, 169–178 (2013) doi:10.1038/ncb2647. In this paper, to identify Rab proteins in ER, first they isolated ER vesicles. Then washed them to remove cytosolic proteins. After washing, these vesicles were solubilized by detergent. Then, it was loaded on to a GTP-agarose column (to bind GTP binding proteins to the column). In the last step GTP binding proteins were eluted with GTP and analysed by SDS-PAGE.
As a control (before loading on to the column) they used GTP gamma S, because it prevents GTP-binding proteins from binding onto the column.
When I looked at their figure; on the control lane, there are lots of bands! So I'm confused :(If it prevents GTP-binding proteins from binding onto the column, after eluting from GTP agarose column why there are bands? And what can these bands be?)

If you can add a link to the article (hopefully it's open-access) that would be helpful in interpreting the results.

The title of this article is ''Rab10 GTPase regulates ER dynamics and morphology''
Link: http://www.nature.com/ncb/journal/v15/n2/full/ncb2647.html
Unfortunately, it is not open-access..

I can still see the figures, though. Which one did you have the question about?

Sort of related: http://biology.stackexchange.com/questions/3258/why-is-gtp-gamma-s-non-hydrolyzable

I had read the answer to the question in this link. But I couldn't understand exactly. So, does GTP-gamma S bind only to the GTPases very strongly? Is this the reason?

This is the figure the question is about. On the right is the control experiment with GTP-γS, on the left without it:

The bands that are visible in both experiments are unspecific binding. If GTP-γS doesn't affect their presence, the mechanism by which they bind to the column can't be specific to the GTPase functionality.
The proteins the authors were after are those that are present in the experiment without GTP-γS, but are not eluted in the experiment with GTP-γS. Those are binding via an GTPase functionality to the GTP on the column. They are marked with arrow heads in the figure.
The general idea behind this control experiment is to distinguish specific binding from unspecific binding to the column. Being able to do that means that the authors had fewer proteins they needed to investigate further, saving them some work.


text:
Is it possible to give a person an allergic reaction in a very specific place?
Can I give a person an allergic reaction at a very specific spot in a tissue?
And if so, how accurate can i get?

You might want to look at the patch test.  Though I'm a little concerned by the motivation of this question.

Thank you this did help :)
Lets say I know a persons allergic reactions will it be possible to do the following? and if so can I change the hypersensitivity level reaction in this proses?

I don't have enough information to make a full answer, but I'm allergic to mangoes. When I eat one, the parts of my skin that the mango touched will get a bad rash, but anywhere it didn't touch will be unaffected.

Depends on the kind of allergic reactions you want to create. If your target tissue is skin epithelium sure you can.
That's actually the way the skin allergy test is all about: The physician tries a wide variety - last time I did it were 28 well known kinds - of substances in liquid form (drops). If an allergic reaction is caused, the physician takes note. Even very small quantities can cause allergic reactions.
These tests are usually made in a secured environment (physician's office, hospital, etc) at a very low level which most of the times, if result is positive will cause just an irritation. The level of irritation will be self-explanatory, so even a non trained patient can understand what is going on.
Not sure if that what you were asking though.
Best Regards,
pa

Funnily, the severity of the reaction does not correlate to symptoms the allergy might cause... of all allergies I have, dust mites gave the weakest skin prick reaction; but it's the only symptomatic allergy I have. The prick test can vary a lot, depending on the purpose; I don't think there's a standard set of allergens they test, apart from ready-made allergen "sets" such as "grasses" or "trees".

Well, yeah I was referring to allergies about trees, grasses, dust, etc.  But sure, allergies are not 100% understood anyway.

Thank you this did help :) Lets say I know a persons allergic reactions will it be possible to do the following? and if so can I change the hypersensitivity level reaction in this proses?

You say "do the following", but there's no "following" sentence, so I don't know what you mean by "do the following". :-) - The hs reaction usually is "linear" but might have more than one variables, so I'd say that it's kind of dangerous to try it out on your own... Better visit a physician.

as in following i mean the patch test mentioned earlier :)
and i wont this is a part of sum recerch Im douing ill go to the universality to figer out more soon thank you.


text:
How to do western analysis to lung tissue?
I am working with lungs from mice, and I want to do a western blot analysis to my samples.
I am having trouble with this because my images turn out very "messy", with a high background.
Perhaps I should filter my samples following their homogenization?
Has anyone had this problem and knows how to avoid it?
What is the best method to homogenize the samples?

Doing westerns with primary tissue can be tough, especially because of the presence (sometimes at high levels) of extracellular matrix (ECM). This material can be quite resistant to homogenization and some lysis buffers. One method I have found to work is to snap-freeze the tissue in liquid nitrogen (not on dry ice or at -80°C) directly after removal from the animal, or after any sub-dissection you may wish to do. Then, when you are ready to prepare your sample, grind it thoroughly with a mortar and pestle in LN2, collect the dust and add a good lysis buffer like RIPA at 2-3X concentration, along with protease and phosphatase inhibitors, also at 2-3X. Then, instead of simply homogenizing, use a probe-tip sonicator (not a water bath one) and sonicate the lysate on ice with 8-10 pulses of perhaps 3-5 seconds each, ensuring that the lysate does not heat up too much, and making sure it does not froth up by getting the tip too close to the surface. Sonication sends very powerful shearing forces through liquid and is excellent at destroying membranes and breaking apart ECM. From here you can do your protein concentration assay (remember to use the same concentration of lysis buffer as your blank), aliquot the lysate and store at -80, or add SDS sample buffer and run your gel.
I routinely use sonication on all of my western samples, regardless of whether it's tissue or cell lines I'm dealing with, and I almost never get smeared lanes anymore. Sonication reduces the viscosity or "thickness" of the lysate, allowing you to pipette it much more easily and accurately. It is also useful when your sample "coagulates" after adding SDS sample buffer, and in this instance it doesn't matter if the sample gets hot, as you'll just be boiling it anyway.
I hope this helps, please let me know if you're still having problems. I used to do troubleshooting like this for a living :)


text:
Why our body does not produce polyunsaturated fatty acids?
Our body does not produce two polyunsaturated fatty acids (PUFA): linoleic acid and alfa-linolenic acid.
I am thinking reasons for it.
Saturated fatty acids have more energy than unsaturated.
Saturated fatty acids do not need NADPH and some many other enzymes as unsaturated in beta oxidation.
This means that less storage places in the body needed to have energy in the form of saturated fatty acids such as for muscles and heart.
There are four main reasons why our body does not use PUFAs as the primary source of the energy but saturated:

PUFAs lower metabolism and interfere with thyroid function
PUFAs spontaneously oxidize, speed up the process of glycation, since too much glucose
PUFAs decrease mitochondrial respiration - more oxygen and CO2, less lactate

where one reason is missing.
One complication of PUFA is

PUFAs promote diabetes, cancer, inflammation and biological stress

which cannot be thought as a reason why our body does not use PUFAs as the primary source of the energy.
There are positive sides of the PUFAs when they are used in other way.
- PUFAs replace trans fats and saturated fats in certain types of foods.
- PUFAs can help your body to eliminate high cholesterol levels
- PUFAs decrease the risk of heart disease
There are also some types of essential fats that your body cannot produce on its own - omega-6 and omega-3 fatty acids.
PUFAs are used to create these.
You can get PUFAs from vegetable oil, fish and nuts for instance.
Balanced diet is essential.
Normal lipid metabolism depends directly on food lipids.
Both the essential fatty acids and right amounts of PUFAs can be obtained from food.
Body converts the essential fatty acids to long PUFAs, which serve as the precursors of prostaglandins and leucotrienes for instance.
There are so many reasons why our body does not produce polyunsaturated fatty acids.
Why our body does not produce polyunsaturated fatty acids?

I think the framing of this question is wrong. I would guess that our body doesn't produce PUFAs because they were present in adequate amounts in our ancestors' diets, so there was little pressure to gain the ability to synthesize them. The common ancestor of animals probably just didn't possess these enzymes, and we've evolved to make do with what's available in the diet.

I am interested on the biochemical sides, not on epidemiological. However, good point to bring the epidemiological sides too.

Can you pleasse tell me that how PUFA lowers metabolism and how they spontaneously oxidize, speed up the process of glycation, since too much glucose.And howw PUFAs decrease mitochondrial respiration - more oxygen and CO2, less lactate and how PUFAs promote diabetes, cancer, inflammation and biological stress???if you elaborate these points it'll be helpful for me...

@HayaDar Thank you for your notice! I am thinking this thread again. I fixed one thing in the body. More explanation about those specific sentences can be found in some Biochemistry textbooks.

The question is unclear for me as it is mostly an inconsistent alaborate on PUFAs in human body. An example of inconsistency: you write there are four reasons andthen  enumerate only three of them. And there is an especially confusing part "There are so many reasons why our body does not produce polyunsaturated fatty acids. Why our body does not produce polyunsaturated fatty acids?" It suggests the elaborate contains answer to your question, thus it should be an answer.

Simply, it’s because our bodies add double bonds in MUFAs by delta 9 desaturase which adds double bonds between delta 9 and delta 10 carbons, hence additional double bonds can be formed between existing double bond and the carboxilc group but cannot be formed between the existing double bond and the omega carbon. As a result, our body cannon synthesize parent omega 3 alpha linolenic acid and parent omega 6 linoleic acid, but can form timnadoneic acid which is an omega 3 PUFA

Welcome. Can you add sources to your claims,preferably links to journal papers? That way people can background read on your material.

Animals, including fish, do not possess any DNA for producing PUFA. No cultured fish can produce PUFA internally. However, human is known to produce milk containing DHA.  The only explanation is all PUFA synthase enzyme are exogenous.  They come from food.  This explains some people are prone to be obese and some aren’t, but the reason is not in genes.
Dinoflagellates are known to have large DNAs Some have DNA for producing important PUFA such as DHA.  No land plant, nor animals have the DNA for such enzyme.  Only explanation is the enzyme comes through the food chain exogenously.  Fortunately, these dinoflagellates are ubiquitous in ocean, and thus all the sea creatures are benefitted from these enzymes.  Modern food production by human however, strips these good enzymes by heat processing and other extraction processes.

Welcome to Biology.SE! Without supporting references only an expert in this area could evaluate the correctness of your answer. Consequently, your answer is much more likely to receive a favorable response if you include supporting references (primary literature is best). In addition, I don't think your connection to obesity is correct and it isn't directly relevant to the original question. ——— Please take the [tour] and then consult the help pages for additional advice on [Answer] effectively and then edit your answer accordingly. Thanks!

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


text:
Where are doxycycline's anti-inflammatory properties derived from?
I have been looking at this antibiotic called doxycycline which is used mainly as an antibiotic. It has however some interesting anti-inflammatory properties. Does anyone know where these anti-inflammatory properties come from, chemically?

See this page for information on how to merge your accounts.

Doxycyline inhibits iNOS and TNF-alpha expression.   It also influences secretion of IL-1beta, IL-8, and TNF-alpha, as well as other inflammatory cytokines.
[These are results collected from multiple systems.]
For appropriate references, see: Leite et al (2011). Anti-inflammatory properties of Doxycycline and Minocycline in experimental models: an in vivo and in vitro comparative study. Inflammopharmocology.
As for the chemistry of how doxycycline accomplishes these things, I'm afraid I don't know the answer, but hopefully, this gives you a place to start.


text:
Is there a comprehensive life science techniques/methods database?
There are so many techniques/methodologies in the life sciences that we can use to interrogate interesting questions. The thing is, most of us are completely unaware of the available methods we can employ. Rather, we go with the techniques we are familiar with or that are popular in our subdomains at the time. But that's pretty limiting.
So I'm wondering... we have databases for everything else... is there one for life sciences techniques/methods? Something like this could be immensely helpful in experimental planning. In particular, I think a comprehensive database would help scientists break outside of their spheres of familiarity and to employ less known (but potentially illuminating) methods to their questions.
I know there are journals that publish protocols and methods, but they are fragmented and don't encompass everything.
Does what I'm looking for exist? If not, how might one go about creating such a tool?

There's Benchfly, which is a video-based protocol library:
http://www.benchfly.com/video-protocols.php
There's also JOVE, which is a peer-reviewed video journal that sometimes covers protocols:
http://www.jove.com/

Thanks. I knew about JOVE but not Benchfly. These looks good. By database, I was thinking something a little more 'smart' in that it would help you select experimental techniques based on various parameters that one might set. JOVE and Benchfly are broken down at a very high level (categories like chemistry or biochemistry) but they don't have much more metadata that would facilitate technique selection. Perhaps a future project for me :)

There is also protocol online, but without tags, only sorted by category. It could nevertheless be interesting.

My personal favourite is OpenWetWare. Think wikipedia for scientific protocols and an open access lab notebook.
There's a problem with this things. Despite the common stereotype of scientist being open and good at sharing, my experience is the opposite. Many laboratories are not good at all in sharing their techniques/secrets. They will share the basic stuff that you can find online, no problem, but that's about it.
Seems they're afraid that everyone's else after them, trying to scoop them. I have suggested OpenWetWare to some other labs and they refused for that same reason. Even though some will use it to find protocols, they don't see a reason to share back.

you can use something like https://www.synbiota.com/ where you can store and share your protocols with collaborators. If you make your project open, then anyone in the world can view your protocols.
The nice thing about this is you can reference actual experiments that made use of the protocol (and literature) and this gives better context and understanding about the protocols...

Looks like it's dead now...


text:
How does Topoisomerase II inhibition affect cancer cells?

Topoisomerase II poisons represent some of the most important and
  widely prescribed anticancer drugs currently in clinical use. These
  drugs encompass a diverse group of natural and synthetic compounds
  that are commonly used to treat a variety of human malignancies. At
  the present time, six topoisomerase II-targeted anticancer agents are
  approved for use in the United States, and additional drugs are
  prescribed elsewhere in the world. These agents all act as traditional
  topoisomerase II poisons and function primarily by inhibiting
  enzyme-mediated DNA ligation.

Briefly describe what TopoII does and how it works. Explain what "inhibiting enzyme-mediated DNA ligation" means.
My thoughts for the first question are Type II topoisomerases cut both strands of the DNA helix simultaneously in order to manage DNA tangles and
supercoils. They use the hydrolysis of ATP. It changes the linking number of circular DNA by ±2. After the ends are
cut, the ends of the DNA are separated and a second DNA duplex is passed through the break. The cut DNA is re-
ligated. This  allows the enzyme to increase or decrease the linking number of a DNA loop by 2 units.  It also
promotes chromosome disentanglement. Inhibiting enzyme-mediated DNA ligation means that it prevents the enzymes are not allowed to ligate. This is
prevented by not allowing the formation of a phosphodiester bond.
Cancer cells grow much more rapidly than most cells in the body. What would be the consequence of "inhibiting enzyme-mediated DNA ligation" and why might this be particularly deleterious in cancer cells?
I am really unsure on this one.
I have looked at the Wikipedia answers to this question and they are wrong.

Hi Abigail, this looks like a homework question so it would be nice if you posted your thoughts on what you think the answer might be before we talk you through the bits you're struggling with.

Hey I am new to the site! I added my thoughts on the question.

That's fine, you're question is already improved with that edit!

First question:
In general, topoisomerases are enzymes that relieve supercoiling in DNA strands and tangling in chromosomes. Type II topoisomerases is special among eukaryotic topoisomerases, considering they use divalent metal ions and ATP hydrolysis in their active site. The enzyme cuts both strands of the DNA helix simultaneously (double strand break) in order to manage DNA tangles and supercoils. It changes the linking number of circular DNA by ±2. After the ends are cut, the ends of the DNA are separated and a second DNA duplex is passed through the break. The cut DNA is re- ligated. This allows the enzyme to increase or decrease the linking number of a DNA loop by 2 units. Inhibiting enzyme-mediated DNA ligation means that it prevents the enzyme to re-ligate the DNA strand after it is cut. In other words, the prevention of a phosphodeister bond between nucleotides.
Second question:
In normal cells, DNA topoisomerase II plays several vital roles, mainly to relieve DNA tangles and coiling. This is done by generating "transient double strand breaks" in the DNA strand, which can sometimes cause harmful translocations and damage. Damage of this type is linked to certain specific types of leukemia. This intrinsically "harmful" quality of DNA topoisomerase II can be exploited in cancer cells due to their high replication rate . Certain cancer drugs, such as etoposide, act by altering the active site of topoisomerase II, preventing (inhibiting) the enzyme from ligating cleaved DNA molecules, causing permanent DNA breaks (and inhibiting replication). The fast replication rate of cancer cells ensures that there are large amounts of the enzyme present in the cell, increasing the effectiveness of any Topoisomerase II poisons. However, this drug can also act against some other types of tissues with large amounts of Topoisomerase II, such as cardiac tissue. Also, cancer cells are constantly in a growing/replication cycle, so they exhibit large amounts of the isoform Topoisomerase IIa, the type that is most likely to produce permanent and detrimental DNA breaks.
Source:
http://www.ncbi.nlm.nih.gov/pmc/articles/PMC2647315/


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Are lactic acid and prolactin rise after exercise connected?
I've been reading about prolactin, a human hormone. Apparently it's levels within the brain can raise after exercise, although the mechanism is not clear to me. I'm interested of there is any connection between lactic acid build up in muscles during anaerobic exercise and prolactin levels in the days following exercise. Both of these sound similar, which may be a coincidence.

Prolactin (PRL) is a pleiotropic hormone. More than 300 different functions have been described for it (Bole-Feysot et al., 1998), although probably the most well known/studied1 is the maturation of the mammary gland during pregnancy and subsequent stimulation of milk production.
One of the fairly well known situations when PRL increases is after an acute stress (Neill, 1970; Seggie and Brown, 1975; Kant et al., 1983). Although the details of this are still a bit foggy, the reason seems to be connected to the anxiolytic properties of PRL (Torner et al., 2001, Donner et al., 2007).
Anaerobic excercise is considered as a stressor, and therefore the PRL response may be important to avoid hyperactivation of the stress pathway. This has been suggested before (Ohiwa et al., 2007) and the rise in PRL may be triggered by lactic acid, as administration of lactic acid alone can stimulate PRL secretion (Sculthe et al., 2013).
Note that this does not exclude that there may be other factors involved in this response.

1 at least in female mammals, but bear in mind that PRL is present in all vertebrates, males and females alike.


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What kind of microscope should be used to view biological structures (such as spores) roughly 5 µm in length?
I would like to be able to identify genera of fungi based on the shape, size, and colour of spores. So the requirements are:

Must be able to view a 5 micrometre object clearly with reasonable resolution
Must be true colour
Must facilitate the measurement of an object within reasonable accuracy (500 nanometres would be nice)
Should be inexpensive (\$100-\$400) fingers crossed
Bonus: Be able to take photos

What type of microscope do I need? What magnification/resolution? Is there a good rule of thumb relating size of object with magnification?
How do I find and purchase a decent microscope after defining my requirements? Any tips?
Blue light is ~450 nm, so does that mean I will not get very good resolution?
(For the sake of context: I have absolutely no formal experience in biology, but I have recently taken an amateur interest in mycology and might enrol in an undergraduate course as part of my degree at some point.)

From Wikipedia:
"With an optical microscope having a high numerical aperture and using oil immersion, the best possible resolution is 200 nm corresponding to a magnification of around 1200×."
So, I searched for 1000x optical microscopes and  found a AmScope 40x-1000x on Amazon. Has anybody used this brand? Will this be sufficient?

Ooh, this one takes photos: OMAX 40X-2000X Digital Binocular Biological Compound Microscope with Built-in 3.0MP USB Camera and Double Layer Mechanical Stage Is OMAX any good?

In this price range, I would avoid anything with very high magnifications > 600x or so. You'll find it very difficult to get enough contrast to see any details.

I think you have a good question, but if you want to get a good understanding of the issues you raise with it, then you really ought to consider spending some time reading this optical microscope primer. In my opinion, you need not bother with taking an undergraduate course at a physical university. As an intro to biology (which is not really necessary if all you want to do is look at small things in a microscope and understand what you're seeing), you might want to consider enrolling in this free class, but if all you want to do is look at small things and have a basic understanding of what you're seeing, I think all you need to do is read the primer.
If it was me, I'd read at least some of the primer before investing the money in a microscope. Otherwise you're just taking a blind leap of faith from someone's (perhaps biased) suggestion.

I agree with Kevin F, if I was you, I'd take a bit of training before buying something or even trying. Biology teachers or staff from the university are usually happy to provide you with a microscope to test something, so just ask. If you want a resolution that high, there are a lot of adjustments to do, commonly referred to as "Köhler illumination", because otherwise you won't get a nice picture.
To answer your question, you can basically forget microscopes at that price range for this resolution, let alone the ability to take photos. You would have to spend >$1000 to get something good enough. As in photography, this is largely due to the lenses. I don't have much experience with fungi, but you might want to have a phase contrast microscope to be able to see the specimens properly. You can buy microscopes with have a port for a DSLR camera, with which I've made good experiences.
For a 5 µm object, you could use a $60 \times$ lens (which, together with a $10 \times$ ocular makes a magnification of $600\times$), or even a $100 \times$ lens, when you want to resolve surface structures, but no idea, if the latter really works with your specimens, so you'd have to test that before buying.
So: First, get a bit of formal training, as suggested by Kevin F. Then, try it yourself without buying, and try it with your own specimens! Then, save money and buy. :-)

Agree 100%. Cheap lenses will just give blurry and distorted images. Beware of promises of 1000x, and color camera for $200... you will just be disappointed. The ideal would be going for something like the Zeiss Primo Star which goes for around 1500-2000€.

I know this thread is old, but I just wanted to say that before reading your suggestion of taking a look at my university's microscopes before buying one I had thought about the same thing and have been just told today by a lady at the university that I would need consent for research from a professor for that.


text:
Where would Saccharomyces cerevisiae be found in highest concentrations in environment?
I have bakers asthma specifically to 'Sc' yeast, and know to avoid bakeries, breweries, etc...
I have figured out to not disturb rotting leaf piles, and fermenting bark mulch. Where else?
What percentage higher are concentrations in say Florida, as opposed to New York state?

interesting question.  aerosol yeast and fungi must vary a log with the weather, humidity and this sort of change in condition.

Here is a link to an interesting article which reports a survey of wild isolates of Saccharomyces cerevisiae in China.

Wang, Qi-Ming, et al. (2012) Surprisingly diverged populations of Saccharomyces cerevisiae in natural environments remote from human activity. Mol Ecol 21: 5404-5417

A relevant quotation from the article:

In addition to grapes and oak tree (Quercus spp.) bark, S. cerevisiae was successfully iso-lated from a variety of damaged fruit collected in orchards or markets in different provinces of China; from the bark of different deciduous trees; forest soil and rotten wood collected in primeval, original secondary and planted forests located in different regions covering temperate, subtropical and tropical climate zones from northern to southern China. Unexpectedly, the success rate of S. cerevisiae isolation from fruit samples (6.5%) was lower than that from tree bark (16.5%), soil (10.8%) and rotten wood (9.2%) samples. S. cerevisiae was more frequently isolated from forest soil samples (success rate 13.7%) than from orchard soil samples (success rate 9.1%). Among the fruits giving positive S. cerevisiae isolation, grape samples showed the lowest success rate.

Table S2 in the supplemental information records isolates as follows:
climatic zone    sites   isolates
__________________________________
tropical           5       20
subtropical        2       14
temperate          13      51

Superficially, from this it looks as if there would probably be little difference between New York state (humid, continental) and Florida (humid, subtropical).


text:
What are the factors to consider when determining the sample size for a given study?
Say I'm designing a retrospective study (where I'd collect data from various schools for the last 5 years) to see if consuming milk can contribute to childhood obesity.  How do I decide on how many schools to visit? If I remember my stats correctly, in general, the goal is to achieve 95% confidence in our results, right?  So how many schools would I have to collect data from so that I will have 95% confidence in the outcome of the study?

Related Question

this is a question of statistical experiment design.  The statistical power of the experiment must include all the factors you are controlling for - age, geographical region, family income etc etc.  So this really depends.  not sure this can be answered here without a lot more information.

You might consider posting this on Cross Validated, but as shigeta mentioned, you'd need to provide more details.

Thanks...actually that's the kind of information I was hoping to hear. The age is 5-10 years, geographical region is US, and family income is irrelevant (randomized).

Maybe this isn't the answer you're looking for... nobody who has ever done a study like this has wanted to have fewer subjects than they could get.  I'd say the answer to your question could just be 'funding'

That's a good point...

I think it depends on what type of analysis you are going to do, but I know for at least an ANOVA you can calculate the smallest sample size for a given power, but in that case you have to have an idea of how big difference you are looking to find-.

About experimental design and is a purely statistical question.

I'm voting to close this question as off-topic because this question is about statistics, not biology.

@Remi.b Agreed, it is also far too broad for here or Cross Validated. There is an entire tag (although unfortunately populated mostly with other poor questions with no answers): https://stats.stackexchange.com/questions/tagged/sample-size

Could be as much as possible – using multistage sampling. But it all depends on how much you are willing to spend, and your time constraint.

Welcome to Biology.SE! This does not seem to answer the question. Please add some more details and references to your answer.


text:
Is it true that oxygen has a diffusion rate in air 10,000 times greater than in water?
According to the Wikipedia page on the design of gills, oxygen has a diffusion rate in air 10,000 times greater than in water. Is that true?
The WP entry refers to: M.B.V. Roberts, Michael Reiss, Grace Monger (2000). Advanced Biology. London, UK: Nelson. pp. 164–165.

There's no clear biological scope in this question, although a good answer has already been posted.

I've slightly rewritten the question to make it clear that the OP was coming at this from a biological perspective. (The OP had already linked to the page on gills and cited the Advanced Biology textbook.)

Yes, according to calculations presented here and here the conclusion is that:

The O2 diffusion coefficient in saturated air (15% oxygen) is 5,700 to 10,800 times greater than in water (60°C and 20°C respectively).

And here is a paper using this difference to investigate the transfer of O2 through the tracheal system of a click beetle.


text:
Does ethanol destroy RNase?
I've gotten conflicting advice on this: some people believe one can remove RNase contamination simply by spraying the bench, pipettes, gloves, etc. with ethanol. Others think ethanol does not destroy RNase and special reagents are needed to prevent RNase contamination.
So does ethanol remove RNase contamination? If not, what reagent does?

See the answer from shigeta below - "inhibition" isn't really the right way to think about this, since that may imply a specific interaction when talking about an enzyme.

Good point, I changed the wording of the question to convey the need for permanent denaturation.

I think the protocols to clean glassware take no chances and just care about RNA degradation rather than targeting one class of enzymes.
EtOH is supposed to denature RNAse and any other proteins on the surface.
Other chemicals such as Diethylpyrocarbonate  (DEPC) used to wash glassware kill all biochemical activity by reacting with nucleophilic moieties in the protein (-OH, -SH -NH(2/3) ).   (See comments below for more.)
These treatments are followed by heating at 300-450 F for 2-4 hours.  This is not a specific RNAse decontamination - all biological activity is killed.
RNAse activity is not assessed by specific detection of a particular enzyme, but rather judged by the degradation of an RNA sample in the lab environment.  So specific inhibition isn't worthwhile - any protein that might interact with and degrade RNA is called an RNAse in these protocols.

In short, RNA contamination and degradation are super easy in a lab setting. We have an RNA-only dedicated workbench in our lab which gets treated with a "broad spectrum" RNAse-degrading solution before any RNA work is conducted.

The heat treatment is called depyrogenation and not only completely removes all biological activity, it also destroys pyrogens (fever inducers) such as LPS or endotoxin.

Thanks for adding to this guys.  I think you can see how the reagents really clean up the glass. I'm familiar with the protocols, but I've not worked with RNA myself.

You don't want to only inhibit or temporarily denature RNAses, if you work with RNA you have to permanently inactivate the RNAses. I work with RNA, and I haven't seen anyone use ethanol to remove RNAses, I would not trust it to work reliably. RNAses are really stable enzymes, they even survive autoclaving to some part, so I would not be suprised if they can refold after ethanol exposure.
The usual methods to remove RNAses are

Treatment with 0.1% DEPC (heat up to at least 60 °C, or autoclave if possible to remove residual DEPC later). Cannot be used with e.g. Tris buffer or anything else that reacts with it. DEPC is carcinogenic, so you should follow the proper safety procedures.
Anything made of metal or glass, heat up to 250 °C for around 2 hours
1M NaOH for at least 30 minutes (you need to wash thoroughly to remove the NaOH)

You should be aware that anything that destroys RNAses will likely do the same to your RNA.
Autoclaving does not protect completely against RNAses, but it still reduces RNAse activity significantly. So while you should not rely on autoclaving to get rid of RNAses, in my experience it is sufficient if you start from a source that is unlikely to contain a lot of RNAses. Pipette tips (not loose tips out of a bag, to minimize handling) and millipore-filtered water are pretty safe after they are autoclaved.
There are also commercial RNAse inhibitors, but I've only used them in cases where I had to mix RNA and protein, and the protein wasn't completely RNAse-free.


text:
Cell Respiration and Oxidative Phosphorylation
Why and how does the proton travelling through the stalked particle in the inner mitochondrial membrane, cause a conformational change in the ATPase?

The passage of protons through the FO (membrane) portion of the molecule (driven by the electrochemical gradient of protons across the respiratory membrane) generates torque at the interface between the a and c subunits. This mean that the ring of 10 c subunits rotates relative to the a subunit. The γ subunit in the stalk rotates along with the ring of c subunits, while the ring of α and β subunits in F1 is constrained to remain static with respect to the a subunit because of the interaction with the δ and b subunits (forming the peripheral stalk). So we have two portions of the molecule the "static" a-b2-δ-α3β3 part and the rotating c10-γ-ε part. The γ subunit rotates within the α3β3 part of F1 and drives a three-step cycle of conformational changes (via intersubunit interactions) which are linked to the phosphorylation of ADP.
There is a useful animation here illustrating the three step catalytic cycle being driven by the rotation of the γ subunit.
(image from Aksimentiev et al. (2004)
Insights into the molecular mechanism of rotation in the FO sector of ATP synthase. Biophys J. 86:1332-44)

For me, ATP synthase is one of the most remarkable features of the cell. Beautiful.


text:
What factors must be taken into account for the maintenance of the immunogenicity of ovalbumin antigen?
What factors must be taken into account for the maintenance of the immunogenicity of ovalbumin antigen?
If the ovalbumin is aggregated, could we inject it for induction of immune reponse?

The preparation of a soluble antigen for injection often includes treatment with an adjuvant which usually creates aggregates of some sort. It is my understanding that particulate antigenic material is preferable because it elicits a better immune response.

Do u know the size of antigens which aggregated by the effect of adjuvant?  If the aggregates' size are around 150 um, do they can activate the immune response? I've read a review article  that suggested that the size of particle (Drug delivery system) should be 1 um for the advantage of to be engulf by phagocytes. It seem like, the antigen aggregate more than 1 um, can not be engulf by the phagocytes ?

Ovalbumin is frequently used in immunology to induce an immune response (see some publications about mice and ovalbumin here.
You usually inject it together with an adjuvans to enhance the reaction. The factors that must be taken into account for its maintainance are the same as for the other antigens, so you should boost this "vaccination" from time to time.


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Why can ATP not be stored in excess?
I just learned that ATP can not be stored in excess and is only made by the body when it is needed. What makes ATP, like glucose and fat is what is stored under the skin or wherever. Now why cant the body just produce ATP and store it for heavy, heavy exercise when it would advantageous for it to be ready rather than it going through the production phase and then used. When I say ATP storage I mean in large amounts.

In a cell, ATP and ADP+P always stay in an equilibrium. If  concentration of ATP is more than appropriate, the equilibrium will be disrupted. To again come in an equilibrium (Le Chatelier's principle), the ATP would break in high speed, will be rapidly converted into ADP+P, and abruptly huge energy will come out that will destroy the cell.

Let's compare ATP, glucose and fatty acids in terms of energy storage.

ATP has a molecular weight of 507 Da
Glucose has a molecular weight of 180 Da, and contains the same amount of energy as 31 ATP molecules
Fatty acids vary in size, but a gram of fat contains about twice as much energy as a gram of glucose (or glycogen)

The difference in energy density is huge, you would need enormous amounts of ATP to replace glucose/glycogen as energy storage mechanism, not to speak of fat. You can't put an arbitrary amount of ATP molecules into a cell, you 'll get into problems due to the osmotic pressure lots of molecules inside the cell would cause. Glucose is stored as glycogen in cells due to this effect, which makes one large glycogen molecule out of lots of glucose molecules.
The energy density difference is even larger if you take into account that ATP and glucose bind water, while fat is stored without surrounding water. The actual difference in energy density of glycogen and fat is around 6 times.
ATP is also not as stable as fat, it can get hydrolized in water. This would be a problem for long-term storage of energy.
You'll find some more details in Albert's "Molecular Biology of the Cell"

I think @AlanBoyd and @MadScientist have touched on the answer, fat is better suited by density for storing energy than ATP; ATP is optimal for quick conversion to bioenergy.  Look at the question in another way: ATP in bioenergy cycle is dynamic - its an energy flux from food and breath to bioenergy.
Biological energy is used at essentially the same rate at which we take it in.   The vast majority is used as soon as its available. If we were to try to store enough ATP for say an hour the costs would be large.
This back of the envelope calculation (see section 3.8) shows that 1 day of ATP is 64.5kg for a 2800 kcal a day energy intake.  Approximately equal to body weight.
Of course ATP is stored in excess - just a few seconds worth though, 8 if you believe competitive cyclists.  ATP is a pretty small quantum of energy... Even an hour storage would add 12 pounds to an adult body weight.  That's a lot. And what advantage would this give us?  We might be able to engage in high energy activities (which use ATP faster than we can make it) for longer.  But up to now it looks as if improvements in efficiency in generating ATP have been adequate for animals to stay competitive.
Look at how evolution has dealt with the other component of bioenergy: oxygen. We can't hold our breath for more than a few minutes. 22 minutes is the current human record; impressive but still not that long.  Oxygen is in plentiful supply brought in from outside and the adaptive costs to storing oxygen internally would simply not justify building up such a capacity for most of us. It seems even that aquatic mammals only hold their breath for a similar amount of time as we can if we practice.  20 minutes for Orcas. Penguins too.  This isn't meant to be a survey of all animals ability to store oxygen, the point being that storage of oxygen has an adaptive cost that is not trivial.

The Weddell seal can manage 80 min.

ATP is also signficantly less stable than other forms of biological storage molecules, such as fat and glycogen. ATP will also slowly hydrolyze by itself when placed in water. The other methods of storage are likely simply more efficient.

Efficiency depends on the purpose.  ATP is for fast transactions, fat is slow.  carbohydrates are in between.  you wouldn't want to have to metabolize sugar to do the 100 meter dash...

Not a complete answer, but a few random thoughts to start off the conversation:
1) There is another molecule that is used as a fast access store, and that is phosphocreatine which can be used to very rapidly rephosphorylate ADP in muscle. In resting muscle it is present at about 5x the level of ATP.
2) Levels of ATP are also used by cells as a regulatory input - in other words the fall in ATP levels with the onset of exercise triggers a response to replenish ATP through e.g. the breakdown of glycogen. In this view it is useful to have a final stage "energy currency" which can act directly as an enzyme substrate and whose level is a sensitive indicator of current energy demand.
3) ATP is also a substrate for RNA polymerase. If ATP was present at vastly higher levels than UTP, CTP and GTP it would probably cause errors in transcription, and might also interfere with the regulatory role of GTP binding proteins, since it would act as a competitor for binding at the GTP binding site.
4) In any case, if ATP was to be maintained at a much higher concentration for rapid use, presumably as soon as it began to be used the ATP generating systems would have to start up to try to replenish the pool. In other words things would really be no different from the way they are, but would simply operate at a higher resting level of ATP.

There is a high rate of ATP dependent processes in the cell such that ATP is immediately used up just after it has been synthesized.
ATP is very unstable in a water environment. It is easily hydrolyzed thus it is not ideal for storage in the very aqueous cellular environment. This makes it ideal instead to have storage sourced of carbon for ATP production.

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