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What causes mutations in regulatory genes?
In detail, what causes mutations in regulatory genes?

This question is just too vague, there is nothing special about regulatory genes with respect to mutations.

nothing causes mutations in any specific genes.  mutations actually occur as the result of random processes and can mutate any given point in the genome.  mutations may be found more often in some genes or regions of the genome more often because they have a positive selective force that gives them more staying power in the gene pool over time.
Its better to think of it this way: mutations happen everywhere, but they stay in the gene pool more often if they do something useful.  Genes are the active portions of the genome so mutations in genes are more likely to stay around.

Not entirely true;multiple well defined mutational processes that have very specific mutational signatures; for example a large subgroup of APOBEC family of cytosine deaminases specifically mutates cytosines that are preceded  by a 5' Thymine and are followed by A/T/C ; Also - the probability of mutations in various regions of the genome can vary by covariates, which is used in the MutSig CV algorithm to estimate mutation frequencies in genes under conditions of null selection.
See http://www.nature.com/nature/journal/v500/n7463/full/nature12477.html for how processes determine mutations.

I wish to add that my point was that non-selective forces can still result in variations in the frequencies of mutations at different genomic loci. Also, some mutational processes are associated with the generation of very specific oncogenic mutations; Recently my group showed that the activity of APOBEC enzymes was strongly associated with a preference for a certain PIK3CA mutation hotspot instead of another that is equally potent, due to the presence of a target sequence motif in that hotspot.
http://bit.ly/1pdgKzS

interesting @AnkurChakravarthy - mind outlining how much more likely such hotspots are than the background here for the record?

OK - in the case of PIK3CA mutations the two hotspots that are equally potent in terms of selection during tumorigenesis generally occur at a 1:1 ratio across cancers overall, but in those tumour types with high APOBEC activity, the correlation between the activity of the enzyme in the genome and a preference for the hotspot containing the sites it recognises was 0.7 (Spearman's Rho) - suggesting that the process was dictating enrichment for mutations in one hotspot over the other, independent of selection.


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What exactly is meant by the expression "differentially expressed"?
As far as I've seen, this expression is almost always used in relation to gene expression profiling. Unfortunately, I have no background in this area. Can someone please explain this in layman terms?

Although each cell of your body essentially contains the same DNA and the same genes, cells in different tissues express (turn on) different genes under different conditions. Measuring differential gene expression involves looking at the amount of expression for a gene (or set of genes) in two contrasting scenarios. The contrast could be across different times, different tissues, different conditions, different related species, etc.
When, you say a gene is "differentially expressed", this is very context-specific. The phrase means nothing by itself, and it is only useful in terms of the applicable contrast. For example, the statement "gene A is differentially expressed" is uninformative, while the statement "gene A is differentially expressed in liver and muscle tissue" is descriptive--it tells you that liver tissues and muscle tissues have a significantly different level of gene A products. Often the terms "up-regulated" and "down-regulated" are also used to provide additional detail. In the context of the previous example, the statement "gene A is up-regulated in muscle tissues" tells you that the level of gene A products is higher in muscle tissues than in liver tissues.

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What is the total number of rounds of cleavage during mammalian embryonic development?
That for frog is 12, but what about mammalian embryos?
I cannot find the exact number anywhere.

I just have an intuation that the number of rounds of cleavage differ between different mammalian, since human seems to have more than 100 rounds, need to verify still from Gilbert.

It's not a totally answerable question, since some types of cells are going to divide more times than others.  But for an estimate, take as a starting proposition that there are 1 trillion cells in the adult human body. [1]  The average weight for a human is 62kg. [2] Average birth weight is about 3.4 kg.  [3]
So that implies roughtly (3.4/62)* 1 trillion = 55 billion cells in a newborn.
You then take the log base 2 of 55 billion, which gives you the exponent you have to hang on 2 in order to get 55 billion, which is about 35.  Then add one for that additional cell division to get from one to two cells == 36 divisions.
Of course I'm just using math, not biology, so your actual reality may vary.  Certainly some cells will reproduce more often than others, maybe cells actually grow in mass instead of dividing (i.e., baby cells might have less mass than adult cells) so the baby-cell-count could be off, lots of possible sources of error.
[1] http://www.nichd.nih.gov/publications/pubs/fragileX/sub3.cfm
[2] http://en.wikipedia.org/wiki/Body_weight#Average_weight_around_the_world
[3] http://en.wikipedia.org/wiki/Infant#Weight

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Why does non-continuous sleep makes you feel tired?
Let's say, hypothetically, that you fell asleep at 10PM, and you were woken up by a noise at 1AM. If you went back to sleep immediately and woke up at 6AM, you will feel tired even though you had slept for 8 hours. However, if you sleep for 8 hours straight , your body will feel rejuvenated. Is it because of REM sleep being cut off and restarting again?

Yes, but I'm guessing. OTOH, you can be well rested after 1 hour, if you know beforehand (=expect) that it will be short. So, it's more a question of psychology, not biology IMHO.

Sleep is not homogenous - there are different phases of sleep - REM, NREM-1 to NREM-4. And these different stages may play different roles in 'rejuvenating' the body (as you put it).
When you first fall asleep, you go into NREM-1 phase, which is the lightest sleep, and progresses into NREM-4, and for the first half of your sleep cycle, this cycles so you go from NREM-1 > NREM-2 > NREM-3 > NREM-4 > NREM-3 > NREM-2 > ....
On the first half of your sleep cycle, the NREM3 and NREM4 phases dominates, only on latter half of your sleep cycle are REM phase observed and progressively dominate.
Depending on how much you are waken up by, once you escape your sleep cycle, you may re-enter back at a different stage, which may mean you are no longer able to have much REM sleep. Subsequently, you will not have the rejuvenation that is associated with REM sleep and you feel tired.
Interesting note: There are people who have tried a method called polyphasic sleep, where they nap/sleep periodically and regularly, instead of sleeping once a day for 8 hours, say. People who promote this says that people do not feel tired because as soon as they nap, they go in REM sleep, and so the amount of REM sleep you get per day will still be the same. I'll let you make your own mind up about this one.


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Why do some fruits have a much wider range of acceptable sizes than others?
Some fruits such as pumpkins can grow to be 100 lbs. Under different conditions, the same variety of pumpkin can produce a 15 lb. fruit. Both plants are healthy, and look the same except for their size difference. If I took two red apple trees and planted them in extreme opposite conditions, the fruit size would be nearly the same. The fruit quantity would be lowered in the tree grown under poor conditions. Why do fruits have such a variable range of acceptable sizes?

Very interesting question, but this can be likened to asking "why are there more than one different animal species?". I hope someone can come up with a nice and specific answer, other than "evolution".

@chm maybe a good answer could address "R vs K selected species"

Some kinds of apples also have wide range of fruit size. The tree in my grandparents garden have fruits varying from 3 to 12 cm in diameter (when they are ripe).

This is also essentially two questions in one. We could be asking "What can explain the size difference in pumpkins", and "Why are all apples of similar size".

It's fine as one question. Each aspect of jmusser poses here informs the other. It's all related.

Pumpkins, squashes in general, grow on vines, while apples grow on trees. Vines are fast growing and trees are not. Zucchini can be quite large; cucumbers, too. Pears, plums, peaches and other tree fruits do have a reduced variation in fruit size. While I do not know the answer to your question, my background in plant biology tells me that this is an important part of the reason. For example, a watermelon or pumpkin rests on the earth, but a tree fruit hangs. If that fruit is too heavy it falls from the ground before it is ripe - before the seeds are fully differentiated that is. So, while "evolution" might be an answer, it is important to discuss what aspects are at play.
I think it has to do with vine vs tree, and how the fruit is attached and hangs/lays.

If you grow a melon on a trellis, it will grow and fall off under its own weight.

grapes grow on vines, but they are closer to apples in acceptable fruit sizes.

but grape bunches come in a wide variety of sizes.

Does that mean if some external support is provided to say an apple, will it grow to a bigger size? Has there any such experiment done?

Bigger pumpkins have more seeds than smaller pumpkins. When they get ideal nutrition the plant tries to make as many seeds as it can to take advantage of the windfall. Many of these vines with big fruit potential grow in compost and rotting material. So they are more opportunistic a pumpkin seed "in the wild" could land in a barren area or in a heap of rotting material-- if it hits the jackpot it meeds to quickly use those nutrients to make more seeds.
PLants with more consistent fruit sizes don't respond as well to "jack pot" levels of rotting material-- they like more consistent nourishment. I think this is another difference.
Also one large pumpkin is a more efficient way to enclose seeds than lots of little pumpkins.

I suspect the answer to this, is that in looking at agricultural fruits you are looking at plants that have been the subject of artificial selection, and the tree vs vine thing is a practical limit, but not the driving factor... for instance a jack fruit grows on tree and is watermelon sized...
I think if you look at something like a pumpkin, you will likely see that pumpkins have 3 traits:

Indeterminate growth ... they will produce as many fruit as the season will let them.
Individual fruit will try to complete growth and ripen before winter
Vines with fewer fruit can devote more resources per fruit, and therefor larger fruit.

Even in apples; if all of the blooms go to fruit the apples will be smaller (commercially apples are thinned to make market acceptable sized fruit): http://homeorchard.ucdavis.edu/8047.pdf
So I think the real issue is less tree vs vine, and more about growth patterns... it may be possible to have a watermelon sized apple, but the tree will have to be sturdy enough to support the fruit: so it isn't a non-issue, but support is a limiting factor.

Well... If I support an apple, it will still mature at the same size.

that is a non sequitur ... if an apple were to grow very much larger it would require more support.

Yes, but is that the only restriction on size variability within a species?

No it is only a limiting factor... apples are highly variable, there is a good segment on them on "The Botany of Desire" (TV show, probably book)... I think you can sort of generalize it as a distance = Rate * Time problem (it really isn't linear, I assume)... Rate is made up of how the plant responds to the level of growth hormones that it produces with the amount of nutrients it can provide, and the time is the time that it is in the part of the growth cycle that it is actually growing.


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Which came first: The Chicken or the Egg?
Has there been any serious scientific inquiries into answering this age old question?

-1 as the attempts at answers show this question needs to be much more constrained to be answerable.

Perhaps the question can be better worded as: "Are there any scientific inquiries to the Chicken or the Egg question?"

This question is invalid but it's such a known question that it's worth putting forth, if only to give an opportunity to point out why it's invalid.

This question is more a philosophical question. The chicken or egg dilemma is not really framed in the proper way according to our understanding of biology today, it's more than 2000 years old.

Since the egg carries new mutations and recombination in each generation, and grow (usually) with the same DNA to the adult chicken/cock, we may assume the egg always comes first. In other words, the theropod lays an egg a little more avian than itself. It IS a philosophical question that, as many others, has perfectly viable biological approaches. I vote to reopen it. We should stop being so closed to "strange" questions.

Most theories about origin of life say that unicellular organisms were first and multicellularity (like chicken) evolved much later, so in that meaning egg (single cell)* was first. Also, almost all chicken ancestors lay eggs (primitive birds, non-avian reptiles, amphibians, fishes and so on...) so eggs existed much earlier than chickens.
*in the meaning of zygote. The development of chicken embryo starts in hen's oviducts, so it's not a single cell at the moment when hen lays it (Gilbert, 2000).

@Hermann_Ingjaldsson Thanks, I've fixed it. Is it better now?

This is a classic case of "it depends on the context."

The Egg: As Marta Cz-C points out, the amniotic eggs that chickens lay were inherited virtually unmodified from their theropod dinosaur ancestors. So when considering the type of eggs, the egg came first.
The Chicken: From a systematic perspective, you must first have an animal that you would call a chicken before it can lay a "chicken" egg. So considering the animal, the chicken.

The question makes the false assumption that chicken and eggs are different life forms.
A life form at stage x is not a different life form than that very life form at stage y.
You might as well ask of which came first, a 7 year old kid or an 8 year old kid.
But they are the same thing and therefore this question is invalid.

Dunno about any serious scientific inquiries into the answer but I always thought the answer is egg. At some point the modern chicken has come into being as the progeny of two pre-modern chickens, however it had to be an egg before it could be a chicken and its parents couldn't have been modern chickens.
However, all that presupposes that you can draw a line in the evolutionary history of the modern (extant) chicken and say this is modern and what goes before is not. I dunno if that really can be done.
Edit: came across this guardian article: http://www.guardian.co.uk/science/2006/may/26/uknews

"Whether chicken eggs preceded chickens hinges on the nature of chicken eggs," said panel member and philosopher of science David Papineau at King's College London.
"I would argue it's a chicken egg if it has a chicken in it. If a kangaroo laid an egg from which an ostrich hatched, that would surely be an ostrich egg, not a kangaroo egg. By this reasoning, the first chicken did indeed come from a chicken egg, even though that egg didn't come from chickens."

And from Prof. Brookfield of the University of Nottingham:

The first chicken must have differed from its parents by some genetic change, perhaps a very subtle one, but one which caused this bird to be the first ever to fulfil our criteria for truly being a chicken; Thus the living organism inside the eggshell would have had the same DNA as the chicken that it would develop into, and thus would itself be a member of the species of chicken

Evolution does not happen over the course of a generation. You are confounding evolution of a species with mutations of a single individual.

That's why I wrote I don't think its possible to draw a line in the evolutionary history and say this is an extant version of a chicken and the previous generation is not.


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Is it plausible that eukaryotic organelles like flagella and cilia are the result of endosymbiosis with spirochetes?
This was a claim by Lynn Margulis explained over at this link.

The sense organs of vertebrates have modified cilia: The rods and cone
  cells of the eye have cilia, and the balance organ in the inner ear is
  lined with sensory cilia. You tilt your head to one side and little
  calcium carbonate stones in your inner ear hit the cilia.  This has
  been known since shortly after electron microscopy came in. Sensory
  cilia did not come from random mutations. They came by acquiring a
  whole genome of a symbiotic bacterium that could already sense light
  or motion. Specifically, I think it was a spirochete [a
  corkscrew-shaped bacterium] that became the cilium.

And why would our bodies incorporate spirochetes as part of our basic functionality?

There are many kinds of spirochetes, and if I’m right, some of them
  are ancestors to the cilia in our cells. Spirochete bacteria are
  already optimized for sensitivity to motion, light, and chemicals… If
  I’m right, the whole system—called the cytoskeletal system—came from
  the incorporation of ancestral spirochetes.

You can find this on Wikipedia: http://en.wikipedia.org/wiki/Evolution_of_flagella#Symbiotic.2Fendosymbiotic.2Fexogenous_models. In lectures we have also been made aware of the fused membrane of cilia with the cell rather than the separate membranes of endosymbionts. There were more (better) reasons but I've forgotten.

Well, since all cells in a multi-cellular organism originally come from the same precursor cells, it’s hard to argue that some of them have endosymbiotic compartments that are missing in others, and notably their precursor.
Notably, there are species which possess both ciliary and rhabdomeric photoreceptor cells, and their precursor is non-ciliary. This is incompatible with Margulis’ endosymbiosis hypothesis.
PZ Myers has a good summary of the relevant papers [1, 2].

[1] D. Arendt, K Tessmar-Raible & al., Ciliary photoreceptors with a vertebrate-type opsin in an invertebrate brain, Science 306, 2004.
[2] D. Arendt, Evolution of eyes and photoreceptor cell types, Int J Dev Biol. 47, 2003.

To play devil's advocate, I'd say that the fact that cilia in modern cells are fully encoded by nuclear genes (and thus that non-ciliary cells can develop cilia) does not entirely rule out the possibility of an endosymbiotic origin: transfer of endosymbiote genes to the nucleus does occur, with e.g. mitochondria having a highly reduced genome, and with some eukaryote species actually possessing mitochondria (or organelles derived from them) that have no DNA at all. That said, there does seem to be other evidence against the endosymbiotic origin of cilia.

@Ilmari You are right, it’s not a definitive proof. But to specify, what I’m arguing isn’t about nuclear genes, it’s that cell development, which does to some extent recapitulate phylogeny, doesn’t show this evolution. The endosymbiosis hypothesis would imply a non-parsimonious evolution on the gene regulatory level, which first induces a loss-of-function and later a gain-of-function in one sister cell type but not the other. The more parsimonious explanation, a single gain of function (again, on the regulatory level) in one cell type, is incompatible with the endosymbiosis hypothesis.


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Are there genes in humans from the common ancestor of all organisms?
How long ago can human genes be traced? Are there any genes that go back unchanged to the beginning of life on Earth? And if so, how many?

Glycolysis seems to be a core pathway of all domains of life: http://biology.stackexchange.com/questions/2934/why-is-glucose-our-primary-source-of-energy

The term to look for is 'ubiquitous' (genes/proteins). Apart from glycolysis and ribosome there is nucleotide synthesis and some (but not all) amino acid synthesis. This list is still not complete.

I very much agree with bitwise's answer. I just want to point out that even in terms of nucleotide sequence there are some extremely conserved genes.
The most highly conserved are ribosomal RNA genes. The image below shows the conservation of 16S rRNAs from archaea, bacteria, and eukaryotes (eukaryotes do not have 16S in their cellular genome but the gene is present in the DNA of mitochondria and chloroplasts). Residues in red are conserved across all domains of life (image source):

A very nice study from 2008 [1] identified tRNA genes as the next most conserved and ABC transporter nucleotide binding domains (NBDs) after that. The alignment below (taken from the paper) shows the conservation of NBDs from various species. Note that they include eukaryotes, archaea and bacteria:
[2]

If you look at orthologous genes in general, without specifying a specific level of sequence identity, then you can find various genes (notably those coding for the enzymes of the glycolysis pathway as mentioned in the comments) that are conserved across most, if not all, cellular life.
So, to answer your question probably yes. At least, there are genes in humans today that can be found in all or almost all other life forms. Whether there were present in LUCA we cannot know for sure, it does seem likely however given how widespread they are today.
Reference

Isenbarger T.A. et al, The most conserved genome segments for life detection on Earth and other planets, Orig Life Evol Biosph. 2008 Dec;38(6):517-33.

Depends on what you mean by unchanged. If you mean a gene that maintained the same nucleotide sequence, then the answer is no.
If you meant a gene that descends from a very ancient gene, i.e. one can find orthologs in very distant species, then the answer is yes. Many of the basic molecular systems are relatively conserved, including several metabolic paths and information processing machinery. This homology is one of the strongest molecular supports for evolution and also one of the reasons that genetic engineering works so nicely.

One thing still missing from the other answers: How many genes? One answer: about 6400, i.e. ~30% of all human genes can be traced back to LUCA.
To come up with this number, I used the eggNOG database of orthologous groups. This database contains clusters of proteins that are probably homologous. So I checked how many human genes are in clusters that also contain bacterial genes. These are 6416 human genes, belonging to 1320 clusters.
Caveats: In some cases, the genes are too diverged and so their homology is missed. It is also possible that a protein evolved in bacteria after LUCA and then got transferred to eukaryotes, and vice versa.


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Given ATP synthase's structure, how can 3.33 protons ultimately synthesize one and only one ATP?
I am familiar with the structure and function of ATP synthase, but one small detail doesn't seem to make sense. It also happens to be a detail that seems very hard to express.

Depending on the organism, there can be from 10 to 14 c-subunits in the c-ring of ATP synthase. For practicality/consistency's sake, let's assume 10.
Every c-subunit is protonated (from the cytoplasmic side of the mitochondrion) and rotated almost 360 degrees within the lipid membrane before it is ultimately deprotonated, which releases a proton into the matrix. Thus, since the c-ring is 10 subunits big (in this case), 10 protons are being introduced to the matrix every 360 degree turn, which mean that 1 proton = a 36 degree turn.
These protons allow for the gamma-subunit to be turned at the inside of the hexamer (of 3 alpha-subunits and 3 catalytic beta-subunits) found below the rotor. The b-subunits are thus located 120 degrees apart. Every 120 degrees precisely, that beta-subunit changes shape, dictated by the rotation of the gamma-subunit, and enters one of the three states that ultimately catalyze one ATP from ADP and inorganic phosphate (since only one of the three states actually forms ATP). Since the gamma-subunit catalyses the formation of 1 ATP every 120 degrees, a full 360 degree rotation would yield 3 ATPs.
This is where things go awry in my mind. Within 360 degrees, 10 protons and 3 ATPs are made. Thus, that makes (10/3) protons per ATP, or 3.33 protons/ATP. When the gamma-subunit receives a proton, it rotates 36 degrees, but ATP formation only happens at precisely 120 degrees, and the only common interval between 36 and 120 is 360 (when 3 as opposed to 1 ATPs are made)!
EDIT: The gamma-subunit must rotate in exactly 120 degree increments, because we observe it do so, as shown here.

More detailed analysis in the presence of lower concentrations of ATP revealed that the γ subunit rotates in 120-degree increments, with each step corresponding to the hydrolysis of a single ATP molecule.

Logically, this shouldn't happen though, since the closest increments of 36 to 120 degrees are 108 and 144.
To summarize: How can 3.33 protons ultimately synthesize one, and only one ATP, knowing that a literal third of a proton doesn't exist (excluding quarks) and that increments of 36 degrees can never add up to exactly 120 degrees?

References

“A Proton Gradient Powers the Synthesis of ATP - Biochemistry - NCBI Bookshelf”, n.d. http://www.ncbi.nlm.nih.gov/books/NBK22388/.

It's probably not that the rotation has to stop on 120° but rather pass 120°? That's the impression I got from the snapshot animation of tagged subunits anyway.

I've added another section to the question which addresses this comment, thanks Armatus.

I think the point is that the rotation of the c subunits is not directly linked to the gamma rotor's movement. As far as I recall, the stator locks the rotor until sufficient energy has built up, then it will let the rotor move 120° in one stroke (kind of like a spring snapping I suppose), stirring the next a or b subunit. BCM has been a while so I'll check up on that and come back with ref tomorrow :)

Never heard that before, sounds interesting. :)

Ok according to my lecture (whose information probably came from Alberts MBotC), the axle rotation is necessary to push the newly formed ATP off the beta units, so it does sound to me like the rotor could stop "in front" of the next b-ATP to be "pushed off" until enough energy accumulates, then flips, pushing it off, and rotating the full 120° to the next b-unit. The article you linked refers to 120° steps when hydrolysing ATP, i.e. the "reverse" rotation.

But there seem to be some not understood discrepancies anyway. My lecturer mentions that plastids' ATP synthase has 14 c-units but appears to require only 2 protons per ATP (6 per turn but 10 units?).

Armatus is right the axle/stalk acts more like a spring than a rigid shaft. It means a single ATP production does not correspond to exactly 120 degrees as the shaft is twisted then snaps into a new position, I always like to think of it working like an old fashioned trip-hammer. Plus there are several variations of ATP synthase that affect things like rigidity of the shaft or whether the rotor can turn backwards. One variation is whether each rotor subunit actually has a slot for a proton or if the subunits are the same size. http://www.sciencedirect.com/science/article/pii/S0014579301023006

@Armatus, remember that not every one of these subunits may actually have a place to slot a proton. They are not all identical, some are just filler. For some bacteria out of 12 subunits only 8 actually have a site for a proton.

I might be taking a bit of a simplistic view on this, but I'm not sure what the confusion is.
The ring of 10 c subunits constitute a motor which uses a proton gradient to drive the rotation of the FoFc synthase.
 (image reference is a nice lecture on ATP synthase).
The motor of c subunits has a 10 fold symmetry.  Each subunit releases a proton as it drives a 36 degree movement of the gamma subunit shaft.  The sixfold symmetry of the ATP FoF1 synthease is completely separate from this, and every 360 degree rotation, three ATPs are generated.  As long as the c motor keeps spinning more ATP is created.
This question is similar to asking why the gear ratios of a bicycle or a car are 3:2 or 5:3.   The symmetry of the synthase and the c subunit are independent except that the torque generated by the c subunit motor needs to be enough to drive the ATP synthase.  If the proton gradient were stronger it could well be 7 protons per full rotation and not 10. Think of the two as gears and you might get the picture?

In a normal chemical reaction you have to have integer numbers. 3 H+ react with 1 ADP and so on. The beauty of this system is that the electrochemical energy of the proton gradient is transferred physically not chemically to the ADP+P to form ATP. This is why you can break the integer rule that dominates most of biochemistry. The flow of H+ deforms the rotor proteins, they later transfer this energy to their substrates, and fuse ADP and phosphate generating the higher energy ATP compound.

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Can I purify polyhydroxyalkanoates by heating the cells extensively?
Traditional methods of purifying polyhydroxyalkanoates (PHAs) and other bioplastics made by bacteria involve washing the cells with harsh chemicals or strong bases.I'm interested in maintaining the spatial position of the PHA globules relative to one another. Therefore, I'm hypothesizing that by heating these cells extensively, I can lyse/degrade the cellular matter and be left with only PHAs, especially given that some PHAs have melting points up to 180 degrees.
Is this approach to purifying PHAs plausible? Has it ever been done?


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Is there a practical upper limit to amount of nucleotides or genes in a transformed plasmid?
I'm currently working on a synthetic biology project which involves working with lots of different parts. I would ultimately like to integrate these genes by transforming a single plasmid. I've heard (casually) that 9-10 foreign genes on a transformed plasmid is often the practical upper limit for these types of experiments, without a real explanation other than "beyond that, it just doesn't work".
Is there a practical upper limit to size of amount of genes in a transformed plasmid, assuming all need to be expressible? What causes this upper limit?

I think the upper limit is more about the size of the plasmid and the efficiency of getting very large constructs into cells, at least in my experience working with mammalian systems. Beyond that you would need to start thinking about other vectors besides plasmids, such as viral delivery systems, Vaccinia vectors, artificial chromosomes, and the like.

I'll most likely be working with E.coli. Putting aside differences in transformation, would there be any reason to suspect differences in functional expression of each gene? (If there is, I've never heard of it. It would seem like genes should get expressed equally in no matter what lengths of nucleotide).

As long as you can get the construct into the cells in one piece without some kind of degradation or shearing, I don't see why they shouldn't get expressed. Most of my experience with E. coli is just using them as a tool to generate plasmids or proteins, and I know that some of their characteristics are quite different than eukaryotes. I was just thinking, though, would a BAC or something similar serve your purpose? 150-350 kb is a heck of a lot bigger than your standard pUC19...

A BAC might work. Thank you for the advice. You should turn your comments into an answer.

From my experience in the mammalian world (and this may apply to bacterial systems as well), it's not so much the number of genes in the plasmid as its actual size. The larger the construct is, the more difficult it will be to get it into your target cells in one piece, without degradation or shearing. Since the transformation efficiency is lower, you are getting fewer whole constructs per cell, so depending on how you've set up your promoters, the overall expression level can be significantly lower. The trouble with splitting your genes amongst two or more plasmids is that each will have different transformation efficiencies, and there will be a certain (perhaps large) number of cells that don't get the full complement of genes, interfering with phenotype analysis. And again, you'll also have to consider differential expression rates.
However, once you get your vector into the cells in one piece, theoretically they should all get expressed at approximately equal levels (assuming identical promoters). It's possible that steric hindrance among multiple transcription complexes may occur - I just don't know enough about bacterial transcription and the effects of circular plasmids to say.
One way to get around many of these factors would be to use a bacterial artificial chromosome or BAC. BACs are 7-10 kb vectors that can have inserts of up to 1 million bp cloned into them and are then electroporated into cells. One of (the many) cool things about them is they control their own duplication and partitioning at cell division, so succeeding generations should have essentially the same copy number as the original. They were heavily used during the Human Genome Project to amplify large sections of DNA for sequencing, but they are also used in many other studies, including synthetic biology. OpenWetWare has a list of some common ones, and NEB sells pBeloBAC11 systems.

Also bear in mind that being based on the F plasmid, BACs are low copy-number plasmids. This means you may have to take care with antibiotic concentrations, and if levels of gene expression are important in your work you shouldn't expect as much as you would get from a standard high copy number plasmid.

@AlanBoyd thanks for the edits - sorry about mitosis, like I said I'm a mammalian guy :)


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Visualizing secondary RNA structure binding to target
I am designing synthetic ribozymes, and need to see the secondary structure that arises from its flanking arms binding to a target mRNA. I've used Mfold as well as IDT's OligoAnalyzer (Hairpin), but these only allow from prediction of secondary structure of the ribozyme folding on itself (which is thermodynamically less stable that 20-40 perfectly complementary nucleotides along the arms).
Is there any software that will allow for simultaneous prediction of secondary structure of two RNA molecules?

there is a software called UNAfold (same website as mFold) which allows nucleic acid hybridization.


text:
Are there known 3- or 4-nucleotide RNA linkers with no RE sites?
Question is rather self-explanatory. I'm looking for a non-cleavable RNA sequence to act as an RNA linker.
I mention 3- or 4-nucleotides only, but making such a sequence with 2 nucleotides is rather easy as seen below, but I've encountered some difficulties in transcription.

GAGGAAGGGAAAGGGGAAAAGGGGGAAAAA...

This also works for just about any random sequence of G's and A's.

Used as a linker, I don't know if the second part of the question, "secondary structure" can be answered.  This could be very dependent on the flanking regions.  Using poly-A is nice for flexibility, but you will get slipping in your transcription.


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Human perception of time depending on age
From what I can tell and what thus far all people with whom I discussed this subject confirmed is that time appears to "accelerate" as we age.
Digging a little, most explanations I found basically reduced this to two reasons:

As we age physically, a time frame of constant length becomes ever smaller in contrast to the time we spent living
As we age socially, we are burdened with an increasing amount of responsibility and thus an increasing influx of information which impairs our perception of the present

To be honest, neither sounds entirely convincing to me because:

In my perception "local time" (short time frames that I don't even bother to measure on the scale of my lifetime) is also accelerating. Just as an example: When I wait for the bus, time goes by reasonably fast as opposed to my childhood tortures of having to wait an eternity for those five minutes to pass.
Even after making a great effort to cut myself off from society and consciously trying to focus on the moment, the perceived speed of time didn't really change. (Although I did have a great time :))

Which leads me to a simple question (and a few corollaries):

Am I just in denial of two perfectly plausible and sufficient explanations, or are there actual biological effects (e.g. changes in brain chemistry) in place, that cause (or at least significantly influence) this?
Is there a mechanism, that "stretches out" time for the young brain so that weight of an immense boredom forces it to benefit from its learning ability, while it "shrinks" time as the brain "matures" and must now act based on what it has learned, which often involves a lot of patience?
If there is such a mechanism, are there any available means to counter it? (not sure I'd really want to, but I'd like to know whether I could)

If I remember correctly it is known that young kids (<4-5 years) cannot distinguish between long and short time frames... I'll look if I can find the proper references about that. I don't know about adult though.\

@back2dos: How far off am I if I paraphrase your question as: "What is a biological/mechanistic explanation of how people learn patience?"

@SteveLianoglou I think it's related, but definitely not the same.  Patience would be more of a meta-phenomenon of how we emotionally deal with the passage of time, but there are issues of perception that are are more fundamental.  See this Nova video(1:40) which touches on this area a little bit.  It's more about whether the brain actually slows down than differentiating between 1 second or 2 seconds, but its indicative that this is an active area of interest.

@jonsca: Thank you, this sounds interesting, however the video is not available in my region :(

@jonsca: However I googled the guy and saw an interesting talk about possibilianism :)

Oh, I wasn't aware it was blocked anywhere, so I'm sorry about that.  I'm sure you could download his references, too.

This is not really a biological answer, but a psychological one:
One important fact to consider is that the perception of time is essentially a recollection of past experience, rather than perception of the present.
Researchers who study autobiographical memory have suggested that part of this effect may be explained by the number of recallable memories during a particular time period. During one's adolescence, one typically has a large number of salient memories, due to the distinctness of events. People often make new friends, move frequently, attend different schools, and have several jobs. As each of these memories is unique, recollection of these (many) memories gives the impression that the time span was large.
In contrast, older adults have fewer unique experiences. They tend to work a single job, and live in a single place, and have set routines which they may follow for years. For this reason, memories are less distinct, and are often blurred together or consolidated. Upon recollection, it seems like time went by quickly because we can't remember what actually happened.
In other words, it can be considered a special case of the availability heuristic: people judge a time span to be longer in which there are more salient/unique events.
Incidentally, (and to at least mention biology), episodic memory has been shown to be neurally distinct from semantic memory in the brain. In particular, a double dissociation has been shown for amnesics who suffer from semantic or episodic memory, but not both.
My apologies for the lack of citations, but a good bit about autobiographical memories can be found in:

Eysenck, M.W., & Keane, M.T. (2010). Cognitive Psychology: A
  Student's Handbook.

You may also be interested in some responses or references to a related question on the Cognitive Science StackExchange:
Perception of time as a function of age

One more thing to note is, when you are 10 years old, the past year represents 1/10th of your lifetime. When you are 80, the past year is only worth 1/80th of your lifetime.

There may be some clues in neurobiology.
A possibility may be that a person's general emotional state may affect their perception of the passage of time, as argued in this article and references within.
Studies of people with damage to their orbitofrontal cortex (prefrontal cortex region) can experience sustained altered emotional states when compared to control samples.
These altered emotional states seemed to affect how they perceived future scenarios (specifically in a simulated gambling context) and the general passage of time compared to controls.
In particular, emotions such as fear and anxiety tended to "speed up" the passage of time whereas positivity and excitement (particularly with regards to future events) lead to a subjective slowing down of time.
Si it may be just that, as kids, we are just more "excited" in general which made for all those seemingly infinite long summers.
Another possibility is that perhaps it is related to the aging process? As we live longer, I can imagine a scenario where our perception of time would alter (slow down?). It would be interesting to study the perception of the passage of time between long-lived and shorter lived populations in this context.

Perception of time can change drastically during an emergency.
When we are younger, much like during an emergency, the brain hasn't activated very many filters for sensory data.  The young have much to learn about the world and more detail is needed for the brain to make appropriate decisions.  Sensory information is recorded in great detail, making time seem to relatively crawl.
As we age, the brain learns to filter out more and more of what it considers inconsequential data.
A good example of these filters at work is the morning commute.  If the same route is driven at roughly the same time every workday, the brain begins to leave out much of the repetitive events and scenery from it's historical record of time.  Given enough repetition, the record of the event will include so little detail in memory as to almost seem not to have occurred at all.  A 30 minute commute will be perceived to have taken very little time -- if it's remembered at all.
As Ferris Bueller so aptly put it, "Life moves pretty fast... You don't stop and look around once in a while, you could miss it."

interesting way of putting it - I think this is very similar to Jeff's answer actually; it is all about how familiar you are with a situation

Maybe the passage of time is perceived as a function of heart rate.
Waiting 5 minutes for a turn on the swings is 300 seconds for a 2 year old and 300 seconds for a 40 year old.  But that same wait is 575 heart beats for the kid, but only 300 heart beats for the adult.

An intriguing thought, because this correlation exists also for adults, although typically when your heart rate rises, there's a lot of things happening in a short time span, so therefore you have a higher awareness of the individual moment.

OTOH this would mean that endurance athletes would perceive their lives as very short :D

Not necessarily. Although endurance athletes would have raised heartbeats during exercise, they would have, in general, lower than the mean resting heartrates.

First:
As a child, it could be that the brain is not encoded with a lot of experience. Thus, processing of new information is less complex, and conclusions are made much faster. These conclusions are then encoded into the brain, making future experiences more complex.
A child may have concluded everything it is able to conclude within the first few seconds of arriving at that bus stop.
Second:
A brain with a lot of previous experiences, is able to occupy itself with "trivialities", such as watching snow flakes fall from the sky or thinking about what to have for dinner. Visualizing different tastes and smells.
A grown individual might be slower to conclude, taking more factors into account. It may also have more to ponder on.
Third:
The brain is thinking all the time. It is unconsciously cross referencing every sound, smell and other sense with previous experiences and actually quite busy. If there is a limit to the amount of energy that goes into the brain for processing stuff, a larger and larger percentage of those electrons will be busy "retriggering" signals. Almost like a city road map. It will take a short time for a number of cars to travel every possible route in a small city, than in a big city. Possibly, this has to happen in the brain as well, to refresh old memories.


text:
What would cause bands to appear lower on a nonreduced SDS-PAGE gel?
I have a protein gel and I'm comparing the reduced with the non-reduced gel. On the reduced gel, I have a few shorter bands but these bands appear with low intensity. In the non-reduced gel, I'm seeing more of the shorter product? What would case the protein to run "smaller" in the non-reduced conditions?

By reduced and non-reduced gel, do you mean native PAGE under reducing and non-reducing conditions? Does your protein have any cysteins and is it known to form disulfide bridges?

SDS-Page and yes.

So you're trying to run a denaturing gel? Strange, I've only seen protocols under reducing conditions for them, a denaturing gel under oxidizing conditions where disulfide bridges can form doesn't really make much sense.

You just remove the reducing agent. Its a pretty standard procedure and a quick search for non-reducing SDS-PAGE seems to show it.

The major effect of changing the redox environment of a protein is the formation or breakage of disulfide bonds. Under sufficiently reducing conditions no disulfide bridges will be present, while under oxidizing conditions your protein will form disulfide bridges if it has the ability to do so.
Disulfide bridges can significantly change the tertiary structure of a protein and change the behaviour of the protein on a gel that way. Generally, disulfide bridges should make the protein more compact and make it run faster on a gel.
It is therefore plausible that your faster band is the protein with disulfide bridges, which will appear to run like a smaller protein as it is more compact. The slower band then is the protein without disulfide bridges which can adopt a more extended conformation.
I'm assuming you're talking about native gels here, as denaturing gels usually contain DTT or beta-mercaptoethanol in the sample buffer to reduce any disulfide bonds and avoid this effect altogether.


text:
How are there alternative initiation codons?
According to wikipedia and the original complete sequence of the K-12 genome, there are multiple non-AUG start codons such as GUG and UUG. How is this possible? I'm particularly curious about the mechanism of translation initiation in the absence of the canonical tRNAfMet.

Actually, the start codon, no matter whether it is AUG or GUG/UGG, always encodes for Met. So the translation is initiated by tRNAfMet (prokaryotic translation). The 30s ribosome subunit binds to the Shine-Dalgarno sequence and then it scans the dowstream mRNA sequence for AUG and the tRNA loaded with Met, which has the CAU anticodon form the most stable interaction. But apparently, only two-bases-interaction between the start codon (GUG, UUG) and the fMet-tRNA anticodon are sufficient for the initiation of translation (1). I will look more into the literature.
1. Initiation of translation in prokaryotes and eukaryotes

Leave it to Marilyn to clarify what everyone else jumbles. I'm seeing some suggestions that there are novel tRNAs with CAC/CAA anticodons. We'll get to the bottom of this.


text:
What would cause proteins to get stuck in the stacking layer of a SDS-Page gel
Typically when proteins aggregate, they will get stuck at the top of the well. However, we're seeing some protein aggregate in the stacking layer even when we're treating the loading volume with DTT.
One peculiar attribute of this experiment is that we're trying to carry out a Cu(I) catalyzed Azide-Alkyne Click rxn. Without the Cu(I), the proteins run normally. However, after Click rxn, we do see some of our expected clicked product, one of our proteins is disappearing into the top band. Hypothesis is that either the copper(I) is changing the migration or oxidative damage from the Cu(I)-> Cu(II) transition is altering the protein.
Returning to the original question, what would cause a protein to stop at the stacking layer vs. at the combs?
[edit]
According to my labmate who was having this problem, the protein was crosslinking with itself to create fairly sizeable polymers. We also saw ladders of the protein with ascending size. Spinning the clicked reaction removed the issue but also resulted at a lost of the protein. It sounds like he wasn't treating with a sufficient amount of DTT to break up the mixture.
This unfortunately still doesn't differentiate between proteins stuck at the combs vs stuck at the stacking layer.

@TomD, we're using pre-casted bis-tris 10% gels. It seems to be stuck at that border.

Sounds like the copper cross linking the protein or creating aggregates that the SDS  buffer can't break up.  add EDTA to your loading buffer before you cook it?

At the interface of the stack gel and resolving gel is a pH change and a change in gel density.  If you feel certain that your protein is not crosslinking so much that it can not enter the gel than I would think about the pH of the sample and the effect of Cu ions on the stacking effect of the stacingk gel.  Is your sample the normal blue color (indication of pH) when you load it (compared to other samples you load)?  Are you overloading the gel?  I have seen overloading cause the effect you describe because the proteins get concentrated to a very high degree before entering the resolving gel.  Could the Cu ions be affecting stacking itself?
Read http://en.wikipedia.org/wiki/SDS-PAGE.  Specifically the section on buffer systems.  You might want to try a different buffer system.
Finally, you could try an acetone precipitation to try to remove the Cu ions and other chemical and see if that helps.

Most of the other proteins are running through the gel as normal. The non-Cu lanes doesn't exhibit this technical issue. Alternatively, I can try prerunning the gel to get a consistent pH environment.


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Structure of RAP Antibodies (Specifically RAP-5)
[EDIT] - Have just found not one but two papers that address my structure problem. However they concern RAP-1A, so I guess my question is now what is the difference in  structure and function of RAP-1A and RAP-5? Does anyone know of X-ray structure analysis being used to examine RAP-5?

original question
I'm a University Physics student writing a mock review article on what to me feels like a very 'un-physicsy' antibody - RAP-5.
Although my knowledge of Biology is pretty poor, research is going quite well. I've found a lot of papers from the 80s talking about conducting immunohisto(/cyto)chemistry experiments, most of them finding that RAP-5 can be used to determine whether a cell has the ras in it, so they are able to measure the percentage of cells that are neoplastic (I believe this means cancerous) and contain ras and the percentage of cells which are normal and contain ras. (the ras-gene being a proto-oncogene which on mutation can result in permanently switched on ras proteins (p21?), which results in proliferation of cells and therefore can cause tumors). This is all very nice but on first glance immunohistochemistry doesn't seem to involve a huge amount of physics (for my bio-physics assignment), apart from using an optical microscope.
I was hoping to be able focus a section of my article on the physical techniques involved in determining the structure of RAP-5. Although there seems to be plenty of literature on uses of RAP-5, I am struggling to find anything on the details on why it is able to be used in such experiments. In other words, I presume that its function is to bind on to an epitode specific to the ras protein (amino acids 10 -17 have popped up a few times) but I don't know if there is any imaging one can do to have a look at the structure and conclude 'yes this is why it binds to ras proteins and not to others'. Is there a technique that is likely to have been used to examine the structure of RAP-5? Is it's tertiary structure likely to be 'Y' shaped like other antibodies? Does it differ in structure from RAP1-4? (this book informs me that RAP1 and RAP2 have 60% sequence identity, but most sources seem to leave out RAP3-to-5, some evening telling me that the RAP family consists of RAP1A/B, RAP2A/B/C and no others!).
Also, if RAP-5 is an antibody, does this mean that it is produced in the body and gets involved in the ras protein signal pathway in order to reduce too much ras expression? (am I right in saying the amount of expression is the amount of protein the ras-gene is producing?) or is it only synthetically produced and used in experiments to measure the amount and location of ras proteins?
Also there seems to be little differentiation between the differences in the functions of each RAP. RAP-5 seems to be used quite a bit in experiments involving Ha-ras - but not exclusively. Do the different RAPs bind to different variants of ras? Ha-ras being the one unique to RAP-5.

I'm not sure is RAP5 is an antibody. Seems to be more of a transcription factor.

Also be careful with your literature search since there is both the Ras related proteins (RAP1A) and the Receptor associated protein domain (RAP/LRPAP)

Also are you sure you're not talking about RAB5c and not RAP?

The confusion that you're facing is because RAP-5 is actually known as RAB5C (GENEID). The ras superfamily (review) is divided into Ras, Rho, Rab, and Rap. But the Rap GTPases are divided only into two categories, RAP1 and RAP2. On the other hand, there are multiple Rab GTPases which include RAB5A and RAB5C.
There are a few crystal structures of both Rab5A and Rab5C in both human and murine forms.
Now, for RAP-5. RAP-5 is a monoclonal murine antibody which has no relationship to the RAP family of proteins. It's probably called RAP because Spandidos and Wilkie weren't thinking clearly when they named their antibodies RAP1-5. In this case, you're probably better off looking at the antibody Y13-259 which is probably called that since its the 259th antibody they tried.
To answer your questions, specifically, very few people have attempted to get a structure since it is an antibody and everyone knows what they look like. Secondly, the antibody is produced in a mouse and added to tissue. And yes, the RAP antibodies bind to different variants of Ras. Also, no one uses RAP-5 since anti-Ras EP1125Y seems to be more popular.
If you still want a crystal structure, I would hunt down the RAP5 patent and look up the peptide sequence. From there, you can look up similar structures in the IMGT database


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How do you knockout an E. coli gene without disrupting the rest of the gene cluster?
I'm familiar with the method to build Keio collection using single-gene deletion with an antibiotic cassette. However, what happens when there are gene clusters or overlapping ORFs and you still only wish to knockout a single gene?

Let's say that we have gene A, followed by gene B so that they overlap with 100 bases. If you want to knock out only gene A, you can do the following:

PCR gene A and clone it into a vector
PCR gene B and clone it into another vector
Insert a antibiotic resistance marker at the overlapping region of gene A and B

now you don't have functional gene A and gene B

Transform your cells only with the vector that contains gene B

Now you have disrupted only gene A but you have a functional gene B.


text:
How do Proteins migrate in MES vs. MOPS
My gels look significantly different in MES (2-(N-morpholino)ethanesulfonic acid) and MOPS (3-(N-morpholino)propanesulfonic acid). That is to be expected. What I don't understand is why the simple presence of the counter-ion significantly changes the velocity of certain proteins.
Is this due to the different pH (MES is usually run at pH 8.0, MOPS is usually run at pH 7.0) or is the charge distribution on the counter-ion responsible for everything?

@leonardo. I feel that you're correct. Now that I'm looking at the chemical structure, I'm surprised that the single carbon difference creates that much difference in the pKa. However, if one is running a gel at the suggested pH for the buffers, the charge ratio should be the same.

these are non denaturing protein gels (No SDS)?

Well, to rationalize everyone's comments, I think @leonardo is right.
This is a denaturing SDA PAGE gel.  The migration of the SDS Micelles which are negatively charged, depends upon the shielding of the solution around it.  The difference in mobility is because the SDS micelles will experience a slightly different field at pH ~6.2 (MES) vs 7.2 (MOPS).
The thought that these have the same charge would be right at exactly the pH corresponding to the pKa.  For these two ions the proportions will not be the same when running 0.8 pH units from their respective pKas. the ion concentration goes as +/- log ([BH]/[B-]) and the charge environment of the buffer should not be the same.  I think that the higher pH for MOPS running will tend to create more negative charge in the solution from MOPS- but also OH- in solution, slowing the mobility of the micelles and favoring the resolution of the larger proteins with MOPS.
This is all given even if the buffering ion concentration is the same.
I'm sure the gory details are buried in the musty tomes of some physical biochemistry journals deep in the library, but this is how it sounds to me.

Congrats, you get my reputation. @Leonardo gets props for showing us the way.


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Ideal low-protein binding membranes
I'm looking through the literature on the topic. It seems like hydrophilic PVDF membranes are ideal for low-protein binding but it also sounds like Regenerated Cellulose is an appropriate substitution.
Looking at one resource (http://www.zellnet.com/elispotplatescolumn/)

When PVDF filter plates were introduced, some investigators chose to use PVDF plates and some continued to use NC.  The reasons for choosing one plate (membrane) over the other are highly varied and won’t be addressed in detail here.  The fact that some laboratories and individual researchers feel strongly that one membrane is superior to the other runs contrary to the large body of Western blotting experience:  Despite some clear-cut differences in how each of the membranes can be used, there is essentially no reported difference in terms of detection sensitivity or signal to noise on NC versus PVDF in the Western Blotting application.  This having been said, it is clear is that the two membranes and their properties are quite different.

The assay that I am looking at in particular is protein/nanoparticle binding to mammalian cells. I am looking for a balance that will enable me to remove free protein without fouling the membrane and/or killing my cells.

May I ask why you need this? Are you a grad student, is this for a high school project, hobby?

@CHM Grad student. I do this for research and the greater good of mankind.

Apparently poly(ethylene oxide) is a biologically interesting polymer, with minimal protein interactions.
The apparently means that I haven't tried it myself, but my polymer teacher told us so.
http://www.sigmaaldrich.com/materials-science/material-science-products.html?TablePage=20204110
http://en.wikipedia.org/wiki/Polyethylene_glycol
Nomenclature seems to favour poly(ethylene glycol). I prefer poly(ethylene oxide), because then I know from what monomer the polymer was synthesized.


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

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

@DanielStandage: why? Biochemistry is definitely on topic

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

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

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

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


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

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

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

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

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

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

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

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

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


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Neutralizing TCA washes
Trichloroacetic Acid (TCA) is commonly used for protein precipitation but the wash waste needs to be neutralized prior to disposal. What exactly is required to effectively neutralize TCA waste? Do I just add caustic?

You might want to add a bit here to establish the biological relevance, as it is now this is a pure chemistry question.

How can I migrate this question?

I've been fortunate to find a protocol hidden away in my lab archives.

The acid waste can be accumulated for 10 days before it needs to be neutralized. The waste should be kept in a carboy in a fume hood.
50 mL 50% w/v Sodium Bicarbonate slurry should be added per liter of TCA waste. During neutralization fumes will be generated which may include chlorine gas.
When the pH reaches 6-8, dispose the waste into the sewer.

I know what the structure of TCA is, which is exactly why I'm asking you how you think Cl2 will be generated?

Why did this get 2 downvotes? Is it incorrect? Why is it accepted instead of the other, much higher scoring answer?

I accepted it since I was the one who asked the question and wrote the answer. It was downvoted since someone decided that it was "unsafe" despite the fact that this is actually the EH&S determined protocol. There needs to be better comment history on SE.

-1. Unless a proper citation is given, I see no reason why trichloroacetic acid would produce chlorine gas in the absence of an oxidising agent.

A neutralization reaction occurs between Brönsted acids and bases to form a salt.
In this case, you have to add a Brönsted base to neutralize your acid. If the final pH is important, you should work out the exact quantity of base to add. Otherwise, just use litmus paper to check the pH of your solution, and add base until it's neutral.
EDIT
In view of the other answer proposed, I want to stress the following:
Do NOT dispose of your solution in the sink/sewer. Leave it in its plastic bin until a lab technician takes it to the furnace.
If the base you use is sodium bicarbonate (NaHCO3), there will be bubbling, caused by the decomposition of carbonic acid (H2CO3) into water (H2O) and carbon dioxide (CO2). You've successfully neutralized the TCA once there's no bubbling upon addition of NaHCO3.
It is indeed best to proceed slowly while adding your base.


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Phagemid display
If I'm using a bacteriophage for phage display and I'm trying to avoid avidity effects by using a helper phage what would be the best way to maintain a large library size while keeping everything monomeric? I'm asking in terms of maintaining an appropriate infection ratio and or phage tricks and tools to maintain the that stoichiometry.

If you don't end up getting an answer here, I would recommend reposting on researchgate

I found article in Nature:
A helper phage to improve single-chain antibody presentation in phage display

Experimental protocol
Standard cloning procedures, determination of colony-forming units and
plaque-forming units, and immunoblot after PAGE were carried out
according to Sambrook et al.
Construction of the packaging cell line.
DH5alpha/pIII
[M13KO7DeltapIII]. The M13 pIII gene fused to promoter P/A1/04/03
(ref. 17) was inserted into the multiple cloning site of the
lambdaattP plasmid pLDR9 (ref. 18). The origin of replication and the
kanamycin resistance gene were cut out and the remaining
lambdaattP-pIII nonreplicative DNA was religated. Escherichia coli
DH5alpha containing plasmid pLDR8 (ref. 18) were transformed with the
lambdaattP-pIII nonreplicative DNA and plated on agar plates
containing 25 mug/ml ampicillin. After incubation overnight at 42°C,
the colonies obtained were replica-plated and ampicillin-resistant
clones were identified, named E. coli DH5alpha/pIII. They were
subsequently transformed with M13KO7DeltapIII, resulting in the
packaging cell line E. coli DH5alpha/pIII [M13KO7DeltapIII].
Production of phage.
To obtain M13KO7DeltapIII helper phage,
M13KO7DeltapIII DNA was electrotransfected into E. coli K802
containing pNDI (ref. 15). To obtain hyperphage, M13KO7DeltapIII DNA
was electrotransfected into E. coli DH5alpha/pIII. Phage were produced
by cultivation in the presence of 0.5 mM
isopropyl-beta-D-thiogalactoside (IPTG) at 30°C for 24 h with shaking
at 200 r.p.m. Escherichia coli Top10F' or E. coli Xl1blue carrying the
pSEXphOx(Yol) phagemid or a scFv library in pSEX81, respectively, were
infected with a helper phage preparation at a OD600 of 0.1 at a
multiplicity of infection of 20 as described.
Phage quantification by ELISA.
Dilution series of phage were coated in
100 mM NaHCO3, pH 8.6 in MaxiSorb ELISA plates (Life Technologies,
Karsruhe, Germany) for 16 h at 4°C. Blocking was done with 2% skim
milk powder in PBS (137 mM NaCl, 3 mM KCl, 8 mM Na2HPO4, 1 mM KH2PO4,
pH 7.3) for 2 h at room temperature. Phage were detected with the
monoclonal mouse antibody B62-FE2 (Progen, Heidelberg, Germany)
specifically binding an epitope on the pVIII major coat protein of the
M13 phage. M13KO7 phage of known colony-forming units were used for
standardization.
Immunoblot.
Approximately 1010 phage were applied per lane on a 10%
polyacrylamide gel. Blocking was done with 2% skim milk powder in PBS
for 2 h at room temperature. Immunostaining was done with the mouse
mAb anti-g3p (MoBiTec, Göttingen, Germany) recognizing the pIII coat
protein of M13KO7, visualized with 3,3', 5,5'-tetramethylbenzidene
(TMB) substrate (Promega, Madison, WI).
Antigen-binding phage ELISA.
Dilution series of BSA-phOx antigen in
100 mM NaHCO3, pH 8.6, were coated in MaxiSorb ELISA plates (Life
Technologies) for 16 h at 4°C. After blocking of unspecific binding
with 2% skim milk powder in PBS for 2 h at room temperature, 2 times
109 single-chain phage diluted in 2% skim milk powder in PBS were
applied to each well for 1 h at room temperature. After washing six
times with 400 mul PBS per well, the bound phage were detected with
mAb B62-FE2 as described above.
Panning.
A human scFv fragment library in pSEX81 with a calculated
maximal complexity of 2 times 107 was generated from peripheral
lymphocyte preparations as described21. Tetanus toxin obtained from
Virotech (Rüsselsheim, Germany) was coated to six ELISA wells (Life
Technologies) at a concentration of 0.1 mug/ml in 100 mM NaHCO3, pH
8.6, for 16 h at 4°C. The wells were blocked with 2% skim milk in PBS for 2 h at room temperature, after which 1011 phage of the library
were applied to each well and incubated at 4°C overnight. After five
washes with PBS/0.1% Tween and five with PBS, bound phage were eluted
with 1 mug/ml trypsin (Life Technologies) in PBS for 15 min at room
temperature. Eluted phage were used for the infection of 20 ml of E.
coli XL1blue at a OD600 of 0.4. Infected bacteria were grown on
Luria–Bertani agar plates containing glucose and ampicillin, scratched
from the plates, and used for the production of antibody phage for the
next round of panning as previously described.

Looks great Devashish Das. I'll wait until I'm sure of not getting a better answer, but I think that paper pretty much covered it.

I appreciate that.

Wow! Thanks for answering such an old question. I'm not entirely sure if this solves the avidity issues though.

@bobthejoe: Thanks!! Can you add a little bit what I am missing in this?


text:
What alternatives are there to the amyloid hypothesis?
Given the recent failure of the Bapi clinical trial, there is a lot of questions that have arised from he amyloid hypothesis. However, I can't really think of many other mechanisms that don't involved plaque formation via protein aggregation.
What are the other dominant proposed mechanisms for the biochemistry of Alzheimers?
Reference: Alzheimer’s Drug Fails Its First Big Clinical Trial. Unfortunately, the results of the study have yet to be presented at a medical conference but will be in September.

Could you link to the relevant studies for those who are not familiar with them?

Well, I am sorry but I fail to understand how that would refute the amyloid hypothesis... it just tells us that a monoclonal Ab against amyloid is not a good cure for Alzheimer's.

@nico, strongly agree but I also get the idea that many alzheimer's researchers are beginning to back off on the mechanism and looking for alternative pathways to attack.

I am not really in that field, so I cannot really comment. However, I would not expect anyone to really say that amyloid plaques are not an important mark of Alzheimer's. They are possibly not the good target for a cure.

Hi all. See this article in Nature - very interesting.

Alzheimer's disease is a very complex field, and I am going to restrict my answer to two particular areas: the neuritic plaques and the neurofibrillary tangles. This area is also of interest to me, hence the protracted answer.
The two pathological hallmarks of Alzheimer's disease, first described by Alois Alzheimer in about 1906, are the extracellular neuritic plaques and the intracellular neurofibrillary tangles which occur (exclusively) in the brain of humans with this disease.
The Neuritic Plaques (Amyloid Precursor Protein and β-Amyoid)
[References to the information given below may be found in Price & Sisodia (1998) and in Lowery et al. (1991)]
The major component of the extracellular neuritic plaques is a small, appromimately 4 kDa protein known as the β-Amyloid (Aβ),  or A4-protein.

The Aβ, whose tertiary structure is that of a β-pleated sheet, is itself derived by proteolysis (see below) from a much larger protein known as the Amyloid Precursor Protein (APP).
A number of isoforms of Aβ are known which contain between 36 and 43 amino acids, the most common of which are  Aβ40 and Aβ42.
The Amyloid Precursor protein (APP) is a very large protein (≥ 695 amino acids) of unknown function which is a normal component of brain. It is predicted to have a single transmembrane span.
A number of isoforms of this protein are also known, which contain  695, 714, 751 & 770 amino acids.
The AA751 and AA770 isoforms contain a domain (Kunitz domain) homologous to a class of protease inhibitors, strongly suggesting the proteolysis plays a key role in normal function, and perhaps in (abberant?)  Aβ formation.
One of the cleavages which produces the Aβ, that catalyzed by γ-secretase, occurs within the transmembrane-spanning domain of APP.  How the protease has access to its substrate within the lipid bilayer is a major source of interest.

Researchers who believe that neuritic plaques and  Aβ hold the key to Alzheimer's disease are known as baptists. Their counterparts, who believe that the neurofibrillary tangles are key, are known as tauists, for reasons that will become obvious (see also here).
For the past number of years, baptists have been to the forefront. In my view, the tauist story is even more interesting, and must surely be important in any final understanding of Alzheimer's disease.
 Neurofibrillary Tangles, Paired Helical Filaments and Tau Protein
Examination of the neurofibrillary tangles under the electron microscope revealed that they had a twisted ribbon-like structure which was called the paired helical filament (PHF) (Kidd, 1964; Wisniewksi  et al., 1984).
PHFs are practically insoluble and for a long time this impared progress.  However, extraction using the detergent sarkosyl, followed by sequence analysis and immunological investigations, produced a  major surprise:

A component of the paired helical filament is the
  microtubule-associated protein tau (Goedert et al., 1988;
  Wischik et al., 1988), a most unusual and interesting soluble protein
  which had already been purified to homogeneity and extensively characterized by Kirschner & co-workers
  (Weingarten et al., 1975; Cleveland  et al., 1977a, 1977b).

A perusal of pubmed shows that at this stage interest in tau increased dramatically!

It is now accepted that microtubule-acssociated protein tau in a
  hyper-phosphorylated state is the major structural component of the
  paired helical filament (Lee et al., 1991; Kosik & Greenberg, 1994).

Some key properties of tau are the following.

Tau was first isolated as  a series of closely related proteins (isoforms) which co-purify with porcine tubulin during successive cycles of polymerization/depolymerization (Weingarten et al., 1975).  On an SDS gel, one sees (with bovine brain tau) 4 bands quite close together from (say) 58 - 64 kDa.
Interaction with tubulin is the only known 'normal' function of tau.  It promotes the polymerization of tubulin into microtubules under polymerization conditions, for example (Cleveland  et al., 1977a, 1977b).
Tau is encoded by a single gene, located on chromosome 17, and differential mRNA splicing gives rise to the brain isoforms (six in humans and four in cow) [Andreadis et al., 1989; Goedert et al., 1989; Himmler, 1989], and a single high molecular weight tau ('big tau') in peripheral tissues (Goedert et al., 1992).
Tau contains a series of imperfect amino-acid repeats, located in the C-terminus of the protein.  These repeats are thought to be responsible for microtubule binding (see, for example, Lee, 1990).
Although tau occurs in a hyper-phosphorylated state in the PHF, and may be phosphorylated in normal brain, phosphorylation is not necessary for formation of paired-helical filaments (Goedert et al., 1996).
The tau gene may be knocked out without any visible alteration of phenotype! (experiments done in mice) (Harada et al., 1994).
Mutations in the tau gene are not known to cause Alzheimer's disease, but are associated with very rare 'taupathies', such as an inherited dementia called frontotemporal dementia and Parkinsonism linked to chromosome 17 (FTDP-17; possibly identical with Pick's disease) [Hutton et al., 1994].
The crystal structure of tau has not been obtained to date.  Current thinking is that the soluble form of tau is mainly a random coil (see von Bergen et al., 2006).

The fact that tau can be knocked  without serious consequence, together with the lack of mutations causing Alzheimer's,  has led to much criticism of the tauist story.
However it still is the major structural component of the paired helical filament (neurofibrillary tangles), and must surely be a key player in the understanding of Alzheimer's disease.

This is not to imply that Aβ and tau are the only shows in town.  As stated above, the field of Alzheimer's disease is very complex
An allele of Apolipoprotein E (ε4 allele) is genetically associated with an increased risk in late-onset Alzheimer's, for example (Corder et al., 1993).
Mutations in the presenilin genes are also associated with the disease (Sherrington et al., 1995).  Presenilins form part of the γ-secretase protease complex involved in Aβ formation (see Takeo et al., 2012).
I do not know enough about these fields to comment any further (although this does not usually stop me).
It should be emphasized, however, that most cases of Alzheimer's disease are sporadic, and only a small subset has a genetic component. It is a disease of the brain, and of aging. There is no known cure, no known cause and no reliable (biochemical) pre-mortem test (although a clinic diagnosis may of course be made).

References

Andreadis, A., Brown, W. M. & Kosik, K. S. (1992) Structure and novel exons of the human tau gene. Biochemistry, 31,  10626-10633.
von Bergen, M, Barghorn, S, Jeganathan, S, Mandelkow, E.M., Mandelkow, E.  (2006) Spectroscopic approaches to the conformation of tau protein in solution and in paired helical filaments.  Neurodegener. Dis. 3, 197-206.
Cleveland, D. W., Hwo, S.-Y. & Kirschner, M. W. (1977a)  Purification of tau, a microtubule associated protein that induces assembly of microtubules from purified tubulin. J. Mol. Biol. 116, 207-225
Cleveland, D. W., Hwo, S.-Y. & Kirschner, M. W. (1977b)  Physical and chemical properties of purified tau factor and the role of tau in microtubule assembly. J. Mol. Biol. 116,  227-247.
Corder, E.H., Saunders, A.M., Strittmatter, W.J., Schmechel, D.E., Gaskell, P.C. Small, G.W. Roses, A.D., Haines, J. L. & Pericak-Vance, M.A. (1993) Gene dose of apolipoprotein E type 4 allele and the risk of Alzheimer's disease in late onset families. Science 261, 921-923.
Goedert, M., Wischik, C. M., Crowther, R. A., Walker, J. E. & Klug, A. (1988)  Cloning and sequencing of the cDNA encoding a core protein of the paired helical filament of alzheimer disease:  Identification as the microtubule associated protein tau. Proc. Natl. Acad. Sci. USA 85, 4051-4055. [pdf]
Goedert, M., Spillantini, M. G., Jakes, R., Rutherford, D. & Crowther, R. A. (1989) Multiple isoforms of human microtubule associated protein tau:  sequences and localization in neurofibrillary tangles of alzheimer's disease  Neuron 3, 519-526.
Goedert, M., Spillantini, M. G. & Crowther, R. A. (1992) Cloning of a big tau microtubule associated protein characteristic of the peripheral nervous system. Proc. Natl. Acad. Sci. USA 89, 1983-1987. [pdf]
Goedert, M., Jakes, R., Spillantini, M. G, Hasegawa, M., Smith, M.J. & Crowther, R. A. (1996) Assembly of microtubule-associated protein tau into Alzheimer-like filaments induced by sulphated glycosaminoglycans. Nature, 83, 550-553.
Harada, A., Oguchi, K., Okabe, S., Kuno, J., Terada, S.,  Ohshima, T., Sato-Yoshitake, R., Takei, Y., Noda, T. & Hirokawa, N. (1994). Altered microtubule organization in small-calibre axons of mice lacking tau protein. Nature 369,  488-491
Himmler, A. (1989)  Structure of the bovine tau gene:  alternatively spliced transcripts generate a protein family. Molec. Cell. Biol.  9,  1389-1396.[pdf]
Hutton, M., Lendon, C. L., Rizzu, P., Baker, M., Froelich, S., et al. (1998) Association of missense and 5'-splice-site mutations in tau with the inherited dementia FTDP-17. Nature 393, 702-705.
Kidd, M. (1964) Alzheimer's disease - an electron microscopical study. Brain 87,  307-320.
Kosik, K. S. & Greenberg, S. M. (1994)  Tau protein and alzheimer disease. In Alzheimer Disease.  Terry, R. D., Katzmann, R. & Bick, K. L., eds.  pp 335-344.  Raven k. Press, New York.
Lee, G. (1990) Tau protein:  an update on structure and function. Cell Motil. Cytoskel. 15, 199-203
Lee, V. M.-Y., Balin, B. J., Otvos, L. & Trojanowski, J. Q. (1991) A68:  A major subunit of paired helical filaments and derivatized forms of normal tau. Science 251,  675-678.
Lowery, D.E., Pasternackj, J.M., Gonzalez-DeWhitt, P.A., Zurcher-Neely, H.
Tomich, C-S. C., Altman, R.A., Fairbanks, M. B., Heinrikson, R. L.
Younkin, S. G.  & Greenberg, B. D. (1991) Alzheimer’s Amyloid Precursor Protein Produced by Recombinant Baculovirus Expression. Proteolytic Processing And Protease Inhibitory Properties.  J. Biol. Chem. 266, 19842-19850.  pdf
Price, D. L. & Sisodia, S. S. (1998) Mutant genes in familial alzhiemer's disease and transgenic models. Annu. Rev. Neurosci. 21, 479-505.
Sherrington, R, Rogaev, E.I, Liang, Y, Rogaeva, E.A, Levesque, G, Ikeda, M, Chi, H, Lin, C, Li, G, Holman, K, Tsuda T, Mar, L, Foncin, J.F, Bruni, AC, Montes,  M.P, Sorbi, S, Rainero, I, Pinessi, L, Nee, L, Chumakov, I, Pollen, D, Brookes, A, Sanseau, P, Polinsky, RJ, Wasco, W, Da Silva,  H.A, Haines, J.L, Perkicak-Vance, M.A, Tanzi, RE, Roses, A.D, Fraser, PE, Rommens, J.M, St George-Hyslop, P.H. (1995)  Cloning of a gene bearing missense mutations in early-onset familial Alzheimer's disease. Nature, 375, 754-760.
Takeo, K, Watanabe, N, Tomita, T, Iwatsubo, T.  (2012)  Contribution of the γ-Secretase Subunits to the Formation of Catalytic Pore of Presenilin 1 Protein.  J. Biol. Chem. 287, 25834-25843
Weingarten, M. D., Lockwood, A. H., Hwo, S.-Y. & Kirschner, M. W. (1975)  A protein factor essential for microtubule assembly. Proc. Natl. Acad. Sci. USA 72, 1858-1862. [pdf]
Wischik, C. M., Novak, M., Edwards, P. C., Klug, A., Tichelaar, W. & Crowther, R. A. (1988)  Structural characterization of the core of the paired helical filament of alzheimer disease. Proc. Natl. Acad. Sci. USA,   85, 4884-32768. [pdf]
Wisniewski, H. M., Merz, P. A. & Iqbal, K. (1984) Ultrastructure of paired helical filaments of Alzheimer's neurofibrillary tangle. J. Neuropathol. Exper. Neurol.,  43,  643-656.

Alzheimer is also considered a tauopathy due to abnormal aggregation of the tau protein. There has been succesful research in that direction, recently, which was a bit neglected, presumably because of vested interests in the amyloid hypothesis.
http://en.wikipedia.org/wiki/Tauopathy
However, another alternative to amyloid (and tau) is the hypothesis that Alzheimer is caused or enhanced by Herpes Simplex virus (HSV-1). One landmark paper found HSV DNA in amyloid plaques and another hint is the association of a lysin-poor/arginine-rich diet with Alzheimer risk (the connection being that HSV thrives in Arg-rich tissue). Also, a cohort study found a direct association.
N. Zilka, Z. Kazmerova, S. Jadhav, P. Neradil, A. Madari, D. Obetkova, O. Bugos, M. Novak: Who fans the flames of Alzheimer's disease brains? Misfolded tau on the crossroad of neurodegenerative and inflammatory pathways. In: Journal of neuroinflammation. Vol.9, 2012, p. 47 doi:10.1186/1742-2094-9-47. PMID 22397366. PMC 3334709. (Review).
A. M. Geppert: [Alzheimer's disease and HSV-1 infection]. In: Neurologia i neurochirurgia polska. 40, 1, 2006 Jan-Feb, 57–61. PMID 16463223. (Review).
M. A. Wozniak, A. P. Mee, R. F. Itzhaki: Herpes simplex virus type 1 DNA is located within Alzheimer's disease amyloid plaques. In: The Journal of pathology. 217, 1, Jan 2009, 131–138. doi:10.1002/path.2449. PMID 18973185.
Letenneur, L; Pérès, K, Fleury, H, Garrigue, I, Barberger-Gateau, P, Helmer, C, Orgogozo, JM, Gauthier, S, Dartigues, JF (2008): Seropositivity to herpes simplex virus antibodies and risk of Alzheimer's disease: a population-based cohort study." In: PLoS ONE 3 (11): e3637. doi:10.1371/journal.pone.0003637. PMC 2572852. PMID 18982063.

Whilst there are several competing ideas for the cause of Alzheimer's, it seems to me that the deposition of β-amyloid (Aβ) is key to the understanding of the disease progression. In many of the papers I will go on to cite, they suggest that prevention of Alzheimer's may be the only cure, as neurons (once damaged) cannot be replaced.
During aging there is a progressive increase in the systemic pro-inflammatory state (sometimes called 'inflammaging'). This is also true for the brain (commonly known as neuroinflammation), and is known to contribute to neurodegenerative diseases by promoting the destruction of local tissues (i.e. neurons) [ref].
Under normal circumstances microglia (resident macrophages of the brain) clear any excess protein depositions (plaques), such as Aβ, in much the same way as macrophages respond to carotid plaques (as can occur during cardiovascular disease). With age, the immune system becomes less functional - a process known as immunosenescence - which contributes to the pro-inflammatory state of aging [ref].
As the immune system becomes less functional, the clearance of Aβ deposits is reduced, possibly contributing to the increased plaque deposition [ref.1][ref.2]. There is also discussion on whether it is the decreased effectiveness of the blood-brain barrier that allows peripheral (from the blood) immune cells to infiltrate the central nervous system, possibly kick-starting the degeneration [ref].
There are many more factors than this, but this is certainly one that interests me as I can see that individuals will have different responses and predispositions to plaques formation in the first place (i.e. the ApoE risk allele, exposure to certain pathogens over the lifetime).
I therefore don't think the study rules out the amyloid hypothesis, in that removal Aβ does not restore the lost neurons (did they really expect it to?). What it does show is that the emphasis should be on prevention, not cure, as 'early-warning' signs, such as biomarkers of Aβ deposits, will be the best way to prevent the loss of neurons.


text:
What is an efficient way to spike 10-50 cells to a culture
I'm trying to add on the order of 10-50 mammalian cells to a mixture of other cells (order of 1e6 cells). What is the best way to do this?
(edit) I guess this wasn't clear. I would really like to emphasize how to sort 10-50 cells to give me an precise (not necessarily accurate but that would be nice too) method of getting a desired number of cells.

If you are you working with primary cell cultures or cell lines may be you could do this when cells are in suspension, after tripsinization, mix them well with a 5mL serologial pipete in a falcon 50mL tube and plate to a 100mm2 with a final volume of 7mL. And finaly draw an eight with the plate in the hood more than 3 times to mix them well.
Edit: For the addition of 10-50 cells, count these cells with trypan blue, and do a 1:100 dilution, for example if you have 1e6 cells in 10mL you could take 0.01 mL (10 microliters) to get 1, 000 cells

And then you add the 0.01mL into 1mL of cell media to get 1 cell per 1 microliter, and then if you wanna get 10 cells, take 10 microliters, and so on

So what about the getting the 50 cells part?

Check the edition of the answer

How do you avoid stochastic effects?

You can't avoid random effects in biology, because biological variability, do a power analysis to ensure you use the correct numer of samples or repeats in your experiment http://biology.stackexchange.com/questions/963/is-it-necessary-to-conduct-a-power-analysis-before-beginning-an-experiment. My answer is a standard method to work with cell cultures, you can also use cell sorting,  to be more precise on your cell count. And try bioforum (a community of methods), to get another solutions


text:
What temperature should mammalian B-Cells be stored at outside of the incubator?
I'm working with murine B-cells. The general protocol is to keep cells on ice to keep them from dying but I've noticed that it makes these cells aggregate and precipitate out. I've heard suggestions that these cells should just be kept at room temperature. How would I be able to determine which conditions at which I should be keeping my cells?
I was trying to collect cells to prepare them for cell binding studies to antibodies and then assess the cells on a filterplate. Typically, you keep them on ice to pause the metabolic state as well as to prevent endocytosis of the bound molecules. However, that shift seems to cause settling. How do you troubleshoot the appropriate temperature to process cells and what are the usual scientific justification for making such decisions.

I know I'm not supposed to ask for clarification here, but I think it's completely necessary in order to provide an appropriate answer. Could you please clarify what you're doing with your cells? i.e. do you need to keep them physiologically active? How long are they out of the incubator? If cells are just going to FACS, ice is fine, just pulse the pellet prior to reading, a little EDTA can also help keeping them from clumping. In general, I would recommend ice or 37 degrees most of the time.

@Rhill45, really?  would say that part of the answer does involve "how long can you keep cell on ice". For instance, if you're going to FACS, what processing steps can you do before running them through the machine, what is the rate of aggregration, etc

Also, why are people collapsing questions from 2 years ago?

For the settling there's little you can do except agitation and using a taller thinner tube and larger suspending volume.
For the clumping however if the analysis permits it, adding serum to a [final] of 5% will help slow the clumping (in the short term). You can also filter out initial clumps by pushing the cells through a nylon mesh filter (they make these especially for flow-commonly used with fixed cells). I've analyzed cells on ice in dark probably as long as 6 hours after harvesting with little change in viability, however this is of course bad practice.


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

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

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

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

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

nice answer Gergana!

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

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


text:
What is the fastest way to build an alanine scanning library?
For interfacial studies, I would like to build an alanine scanning library for one of my proteins examining 20 sites. I will ultimately express the gene using E.coli cell-free protein synthesis. I already have the template gene which was originally built using PCR assembly. Now, what's the best way to build a scanning library?

do you mean, how do i decide which amino acids to include in the alanine scan or how do i create the library of alanine mutants?

@shigeta, definitely more "how".

http://www.genscript.com/alanine_scanning.html  genscript has some pages to help you reason through the design.  Myself I'm not sure, but it looks like it takes quite a lot of oligos to say the least...

Are you still interested in the answer of this question?

@GerganaVandova, not really but it might be of interest to the broader community.

The fastest will change as time passes and better technologies are developed.
I think the fastest method existing at the moment is Shotgun Mutagenesis (provided by Integral Molecular Inc).
This does not employ any new method of doing that. They just provide a set of plasmids, that has all the possible mutations. The set itself is generated by automated DNA synthesis.
So if you don't have a DNA synthesizer with you then simply order the kit from the company.


text:
What are key factors when evaluating and comparing miniprep columns?
I'm looking to comparing different protocols for minipreps for plasmid DNA purification. What factors should I be looking at? A few things come to mind:

Cost
Yield
Time per step
Replacement with other reagents
Spin vs. vacuum
Color and caps or lack thereof

Shopping advice is generally off-topic on all SE sites, and a question about the current price of columns is very localized in time and geographically.

I think this is a perfectly valid question.

Storage conditions and shelf life are some of the things that you should consider in comparing columns.

Seems like you have covered the essentials here. I can't think of anything else.
Since the Qiagen patent on spin prep columns ran out, these kits are very cheap - $0.40 each?  In the 3 or 4 kits Ive used, they all seem to use the same protocol and about the same buffers, so there might be differences in quality or yield but if so, they are small.
You can even make your own buffers as the recipes are on open wet ware.  None of the kits i've used has suggested that these buffers can't be used.  They even usually call the buffers by the same names.
So your application would have be pretty discriminating to differentiate.  Even if this were the case, the quality could probably be improved a lot just by washing the column more.  Can you provide more details about how a few percent one way or another might be an issue?

I guess that it really doesn't matter. I guess we need to form a community wiki.

Congrats shigeta on being the first user to reach 5000 reputation!