text:
What is the criticality of the ribosome binding site relative to the start codon in prokaryotic translation?
In prokaryotic translation, how critical for efficient translation is the location of the ribosome binding site, relative to the start codon?
Ideally, it is supposed to be -7b away from the start. How about if it is -9 bases away or even more? Will this have an observable effect on translation?

I found an oldish paper on this topic (from 1994).  Here's a summary:
Determination of the optimal aligned spacing between the Shine-Dalgarno sequence and the translation initiation codon of Escherichia coli mRNAs. by Chen, Bjerknes, Kumar, & Jay. Nucleic Acids Research. (1994)
Experiment
The authors constructed a series of synthetic RBS regions that varied the length separating a synthetic 5-nt Shine-Dalgarno sequence from the start codon.  The regions varied in size between 2 to 17 nt.  They assayed the activity of a downstream enzyme, chloramphenicol acetyltransferase.
Conclusion
The authors concluded that the optimal spacing between a consensus 5-nt Shine-Dalgarno sequence (5'-GAGGT-3') and the start site was 5 nt. Note: this synthetic SD was made of the last 5 nt of a 9 nt SD consensus sequence.  They also tested a synthetic SD made from the first 5 nt (5'-TAAGG-3')of the consensus SD.  In this case they found the optimal distance was 9 nt.
So the optimal distance depends on where your desired SD aligns with the consensus SD sequence, which optimally is 5 nt from the start.  Read on for more details.
Details

the RBS is considered to be large, extending 20bp on either side of a core Shine-Dalgarno (SD) sequence.  These days, I often hear of the RBS spoken of in sizes that are equivalent to the SD.  So in the parlance of the paper, you question is rephrased as "how does distance of the SD from the start codon effect translation?"
the canonical SD sequence referenced in the paper is 5'-UAAGGAGGU-3'.  It is 9 nucleotides long.  Distances between the SD and the start codon are defined as the number of nucleotides separate the 3' Uracil of the SD from the Adenine of the start AUG.
Example: the distance is 5 nt in the following mRNA
       5'.....UAAGGAGGUnnnnnAUG......3'

if the SD is not a complete 9 nt long, the distance is between the position of where the canonical Uracil would occur.  In the following example, the distance is still calculated as 5 nt:
       5'.....UAAGGAnnnnnnnnAUG......3'

the average distance between the SD sequence and the start codon varies considerably and on average is 7 nt. Other investigators have found "optimal" spacing (circa 1994) ranged from 5 to 13 nt.
the SD site complements with region on the 16s rRNA. The start codon complements to the anticodon of fMet-tRNA loaded into the ribosomal P-site.  So there are two distinct sites on the ribosome that contact the mRNA during translation initiation.

Also good to read

Designing Genes for Successful Protein Expression by Welch et al. Methods in Enzymology, Vol 498. (2011) (pdf)
Automated design of synthetic ribosome binding sites to control protein expression by Salis, Mirsky & Voigt. Nature Biotechnology. (2009). (pdf)

Note
I enjoyed researching this question because I found my own knowledge to be lacking solid empirical details. I do not have any direct experience besides this little lit review with this topic.

Thanks for your thorough answer. I guess I haven't realized that the RBS end is determined by this U base. Does this mean that it can be 100 bases away from ATG? I am actually asking this question, because I have a synthetic construct with RBS annotated on it and some of them are really far away from the start codon. Which could mean either wrong annotation of the following gene sequence, or that the distance doesn't matter that much. But on the other hand I thought that the ribosome should bind both the RBS and the start codon at the same time which I don't see possible with long sequences.

What I am also concerned about, are secondary structures formed between the RBS and the upsream or dowstream sequence. Any thoughts on that? Thanks again :-)

The main idea in the Chen paper I referenced was that the distance between the Shine-Dalgarno sequence / RBS and the start codon is limited by the physical distance between the complementary sites on the RBS. This distance seems to be about 5bp.  According to this old paper, the ribosome should not be able to bind effectively at all if the RBS is farther than say 20bp from the start site because the ribosome can't stretch out far enough to reach both sites at the same time.  New research may have more to say on this topic.

Secondary structure has significant effect on translation efficiency.  Many riboregulators have been observed and designed. For instance, one can design stem-loop secondary structure around the RBS region.  The region will self-complement in the absence of a particular aptamer or a strong complement.  In their presence, it unfolds and allows the ribosome access to the RBS and start site.  Engineered riboregulators enable post-transcriptional control of gene expression. Isaccs et al, Nature, 2004.

It was an interesting question to investigate and I enjoyed learning about it.  I hope more well-posed and provocative questions are asked here and that users react to them by actually going out and reading up on the primary literature.  Looks like that happens occasionally.  Cool.

Yeah, we will see where this site is going...I am also trying the RBS calculator :-)It would be nice if we chat more. Would you write me on gvandova@stanford.edu?

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text:
How is RNAse contamination in RNA based experiments prevented?
Does anyone have any suggestions to prevent RNAse contamination when working with RNA?
I tend to have issues with degradation regardless of whether I use DEPC treated / RNAse free water and filtered pipette tips.

Are you working with isolated RNA or with in vitro transcribed RNA?

@MadScientist: Isolated RNA with a Qiagen RNeasy Kit.

My answer is about avoiding RNAses while working with RNA, I'm not working with isolated RNA so there are probably additional steps you should take to avoid RNAses present during the isolation.

@MadScientist: I still think your answer is very relevant, I could be getting degradation during the down-stream experiments as well.  I was hoping that this question could be relatively general because I know a lot of different people have different processes to avoid RNAse contamination.

You need to be careful about everything that comes in contact with your samples. Every spatula, beaker and every magnetic stir bar need to be RNAse-free.
For metal- and glassware we usually put everything into a drying closet at 200-250 °C for a few hours. That should get rid of RNAses pretty efficiently.
The RNA chemicals should be seperated from other chemicals if not everyone in the lab is working with RNA. They shouldn't be weighed using spatulas, but just poured into an RNAse-free container to avoid contaminating the storage container.
Use disposable plastic wherever possible, you should be able to perform most stuff in falcons and Eppendorf tubes.
Be careful with performing e.g. a DNA-prep at the same bench as the RNA work. The first buffer for the DNA-prep contains RNAse.
Avoid keeping your Eppendorf tubes or falcons open for long, that should reduce the potential for contamination.
We've had pretty good experience with just using water from a Millipore system and autoclave it once, that seems to get rid of the RNAses. I know that this is not a sure thing as the RNAses can survive autoclaving, but it seems to work well enough.

Here are some tips from what I routinely do:

wipe all the surfaces (including pipettes and gloves) with RNAse Away or similar
do everything on ice
use a RNAse/DNAse free tubes. Use also commercial RNAse free water (it's worth the investment).
keep all your RNA reagents separate from your cell culture/animal work reagents. Autoclave all your buffers and don't let them sit forever. Just make fresh buffers every now and then.
don't talk over the tubes

If you keep having degradation following all the above, consider how you prepare your sample. Do you have enough RNAse inhibitors in there? Do you use a RNA-conserving solution? Is your tissue sitting for a long time before the lysis? If your extracting RNA from cells, try using trizol or a similar reagent to preserve the RNA right from the lysis. If your getting the RNA from tissue samples, try snap freezing them in liquid nitrogen.

Apart from tips already mentioned, change your gloves every now and then.
Also, in my experience (with the Qiagen kit), keeping samples on ice shouldn't make much of a difference.

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text:
Are lymphocyte sizes clustered in two groups?
Tortora writes in Principles of Anatomy and Physiology:

Lymphocytes may be as small as 6–9 μm in diameter or as large as 10–14 μm in diameter.

Those ranges are quite close to each others. Should the above be taken to mean that lymphocytes sizes are clustered in two groups, or is it just a way of saying that lymphocytes are 6-14 μm?

Unlike erythrocytes that have a very rigid shape and almost cannot change their size (hence the size distribution is indicative and can be used for diagnostic purposes in medicine), lymphocytes can change their size in a wider range, this is why you see the numbers 6-9 and 10-15 μm.
And they indeed cluster into several different groups: so-called "large granular lymphocytes", also known as NK-cells or "natural killers" (usually >10μm) and "small granular lymphocytes", constituted by a large family of T- and B-lymphocites(usually <10μm). But this clustering is not really distinct, for as I said above, both NK and T/B-cells can shrink (if the osmolarity of the external medium grows due to acidosis, inflammation etc.) and swell (on binding many IgE/IgG complexes, on certain cell factor released etc.). Besides, there are also some intermediate size cells, called NKT-cells, that also flatten the distribution.
So, what you can definitely say is that the typical size distribution of lymphocytes has two peaks: around 8 and 12 μm respectively.


text:
How long does antibiotic-dosed LB maintain good selection?
Various people in our lab will prepare a liter or so of LB, add kanamycin to 25-37 mg/L for selection, and store it at 4 °C for minipreps or other small cultures (where dosing straight LB with a 1000X stock is troublesome).  Some think using it after more than a week is dubious, but we routinely use kan plates that are 1-2 months old with no ill effect.
How long can LB with antibiotic such as kanamycin, chloramphenicol, or ampicillin be stored at 4 °C and maintain selection?

This Bitesize Bio article is very informative on this issue.  The 1970 study they cite found negligible reduction in the the efficacy of non-beta-lactam antibiotics (kanamycin, chloramphenicol) over spans of 4 weeks or 60 days.  Ampicillin loses about 10% activity at 4 weeks.
In my lab, we only take care not to use LB/Amp plates that are older then 4 weeks, for LB/Km, we store them at 4 °C for more then a month, sealed in a plastic bag.

As far as I know, that is fine.  Thats the temperature that Amp is regularly stored at.  I always do a negative control anyways to ensure that my antibiotics are still lethal.  Just don't stick it in the microwave to make plates

Ampicillin degrades by hydrolysis (albeit very slowly if not catalyzed), difficult when in dry, much easier when in solution.

"That's the temperature that Amp is regularly stored at" - and in fact, Amp does lose activity (a few % per month?) as a consequence.

I don't know about solutions, but we keep LB-Amp/Carb/Gm plates in 4°C for two-three weeks at a time, and the antibiotics seem to work.


text:
Is exon order always preserved in splicing?
Are there any cases in which the splicing machinery constructs an mRNA in which the exons are not in the 5' -> 3' genomic order? I'm interested any such cases, whether they involve constitutive or alternative splicing.

When you say order do you mean something like Exon 1 - Exon 3 - Exon 2 - Exon 4? or do you mean events such as exon skipping?

Exon skipping events preserve exon order. So...yeah, I mean something like your first example.

While this may not be what you are asking, given an exon sequence E1E2E3E4, if E2 and E4 have the exact same sequence, it will appear as if E1E2E3 and E1E3E2 can be generated from this gene.

I don't have any literature to back this up but I doubt that it occurs (at least frequently).
For example, imagine a simple three exon gene.  Upon splicing exon 1 to exon 3, exon 2 would be excised as part of the intron lariat and subsequently degraded.  So in order for exon 2 to be spliced to exon three you would need to either have splicing between exon 3 and exon 2 in the lariat or another copy of the pre-mRNA.  This is typically called trans-splicing but it only occurs in specialized systems such as spliced-leader sequences in C. elegans.

After performing a quick literature search, I am, as with GWW, unable to provide any literature against this occurring, although this paper by Black (2005) states that exons in multi-exon pre-mRNAs are always maintained in order.
 Black DL. 2005. A simple answer for a splicing conundrum. Proceedings of the National Academy of Sciences of the United States of America 102: 4927–8..
Separate components of the spliceosome recognise sequences at the the start and end of each intron, along with a branch point adenine and some other conserved (highly in the case of yeast) regions upstream of the 3' end. The components seem to assemble in a particular order, which is 5' to 3' directional.
The general consensus seems to be that the spliceosome moves along the pre-mRNA from 5' to 3' removing the introns and splicing the exons together to form the mature transcript, ready for translation. The introns are subsequently degraded. In the case of eukaryotes some processing is required to transport the mRNA out of the nucleus to the ribosomes in the cytoplasm.
The mRNA would have a start codon near the 5' terminus and stop codon near the 3' end, at the boundaries of the CDS, and it isn't clear to me how this structure could be maintained if exon order didn't follow the 5' to 3' assemblage. I should imagine that the protein could be affected in much a similar way to exon skipping, possibly resulting in a truncated, inactive or ineffective protein. As protein folding is determined by the interactions between the amino acids, any change in exon order (secondary structure) is likely to result in a different tertiary structure.

There are references in the literature to the phenomenon of "exon scrambling" which seems to be what you are asking about, but the prevailing view is that the evidence for this process, which comes from comparing EST sequences with genome sequences, can be explained by cloning artefacts occurring during EST characterisation. Certainly I agree that there is no evidence for a gene in which the rearrangement of exons is part of the normal pathway for mRNA generation.

Shao et al. (2006) Bioinformatic analysis of exon repetition, exon scrambling and trans-splicing in humans. Bioinformatics 22: 692-698

There is also more recent evidence from deep sequencing for rearrangement of exon order via circular RNAs:

Salzman et al. (2012), Circular RNAs Are the Predominant Transcript Isoform from Hundreds of Human Genes in Diverse Cell Types PLOS ONE 7: e30733   DOI: 10.1371/journal.pone.0030733
Most human pre-mRNAs are spliced into linear molecules that retain the exon order defined by the genomic sequence. By deep sequencing of RNA from a variety of normal and malignant human cells, we found RNA transcripts from many human genes in which the exons were arranged in a non-canonical order. Statistical estimates and biochemical assays provided strong evidence that a substantial fraction of the spliced transcripts from hundreds of genes are circular RNAs. Our results suggest that a non-canonical mode of RNA splicing, resulting in a circular RNA isoform, is a general feature of the gene expression program in human cells.

Incidentally it is perhaps worth mentioning that in the case of the lectin concanavalin A the equivalent effect is produced post-translationally when the protein rearranges.

Carrington et al. (1985) Polypeptide ligation occurs during post-translational modification of concanavalin A. Nature 313:64-67


text:
How can I avoid digesting protein-bound DNA?
I'm interested in sequencing and analyzing the bound DNA, and minimizing the amount of unbound DNA that gets sequenced through digestion.
When digesting protein-bound DNA, is all of the unbound DNA digested? Is there a way to maximize the amount of unbound DNA that is digested?

Well, it depends if you just want to sequence your protein bound fragments.
If that is the case, I suppose you could go for a FAIRE-seq.
Basically, you perform several iterative phenol/chlo extractions, thus separating the bound and unbound fragments.
Then you sequence the unbound fragments that are present in the aqueous phase, and after the mapping, you can deduce the regions that were actually bound to proteins.
I don't know if it is possible to treat your bound fragments (that you isolate after a phenol/chlo extraction in the organic phase) with proteinase K, then perform another phenol/chlo extraction to purify your formerly bound fragments and sequence them.
Could be worth trying.

Are you interested in transcription factor binding (and similar), or in histone modifications?
If you are looking at investigating something with a short binding motif that occurs redundantly across the genome, ChIP-exo works: http://www.cell.com/abstract/S0092-8674(11)01351-1
This technique involves digesting one strand of the DNA up until the bound protein-DNA complex via exonuclease, while leaving enough of a tag on the other end to uniquely identify it's location within the genome. I have no first-hand experience with it, and it's yet to be independently replicated, but it looks fantastic. I don't recall whether they investigated the extent to which off-target DNA was digested by the exonuclease, but theoretically I expect a significant portion would have been eaten up.
If you are interested in ChIP of histone modifications, you can use micrococcal nuclease (MNase) for your sample prep instead of sonication. Again, I have no first-hand experience, but I understand that this enzyme can be used to digest DNA until it reaches the nucleosomal core particle, and there are published ChIP-seq experiments employing this strategy. The MNase would be expected to digest unbound DNA fragments.

This is a difficult experiment.  It's usually done by digesting the DNA under conditions where the binding protein is solidly attached.  DNAse is added, and then after some time, the DNAse is degraded with heat or the addition of an inhibitor.
What you get depends on the length of the DNA protected by the protein and the tenacity by which the protein binds itself to the DNA.

If your aim is to figure out which nucleosides are specifically recognized by the protein then once you narrow down the sequence with one of the above methods, you would have to perform sequential rounds of mutagenesis.


text:
Under what conditions do dendritic spines form?
I'm looking for resources or any information about the formation of dendritic spines and synaptogenesis, especially in relation to how new connections are formed on a daily basis.
Does the electrotonic signalling along the axons and through the spines cause new connections to be made based on some kind of spatial condition (maybe an electrical or chemical attraction), or is there some larger heuristic here?

Spine formation (spinogenesis) is almost certainly due to chemical, rather than electrical, signalling between neurons. Although there are exceptions (gap junctions, for one), most forms of inter-cellular communication are mediated by chemicals released by one cell and detected by another. You are right that the cues for synaptogenesis are probably localized (the "spatial condition"), but I'm willing to bet the farm that those local cues are chemical in nature.
A recent paper from Kwon and Sabatini (2011) shows that local release of the neurotransmitter glutamate is sufficient to cause a functional spine to form. Glutamate receptors on the dendrite detect the glutamate and a spine forms (within seconds). At least under these conditions, the presynaptic machinery isn't required at all! Of course, in a less reduced preparation, electrical activity in the axon will signal the glutamate release from the presynaptic side. Thus, in this case, spine formation is activity-dependent but is mediated by chemical cues.

Please note that the phenomenon Kwon and Sabatini describe only manifests itself in brain slices from very young animals and not at all in brain slices from adults.

Dendritic spines are thought to grow and recede under LTP and LTD, respectively. See (Bosch and Hayoshi 2011) for a review.
From there, much of the synaptogenesis occurs due to surface molecules present both on the dendrite and the presynaptic axon in the growth cone.  Localization and guidance are achieved through gradients of growth factors in the developing nervous system  See (Kolodkin and Tessier-Lavigne 2011) for a review of all of these mechanics.
How this maps back onto the human CNS and thinking/learning/memorizing is still up for debate, but some of these mechanisms must have been preserved in higher species.

References:
Bosch M, Hayashi Y. (2012) Structural plasticity of dendritic spines. Curr Opin Neurobiol.,22(3):383-8. (Epub 2011 Sep 28).
Kolodkin AL, Tessier-Lavigne M. (2011). Mechanisms and molecules of neuronal wiring: a primer. Cold Spring Harb Perspect Biol., 3(6). [DOI]

Although both classes of phenomena are relevant for synaptic and spine dynamics: 1) LTP and LTD are classically thought to require a pre-existing synapse, 2) what happens in development may be different than what happens in adult life.

I think the insightful comments you've made on my answer and the one above really are worthy of a new answer.  I've been out of the neuroscience game for almost 10 years now, so I definitely defer to your expertise!  Thanks for commenting.

Two classes of factors contributing to spinogenesis have been described in the literature, based on whether they can be considered extrinsic or intrinsic to the dendrite (my classification). Here is a shortlist of evidence in favour of either:
A. Extrinsic:

the presence of extracellular glutamate facilitates spine
formation in tissue from very young mice (Richards et al., 2005;
Kwon and Sabatini, 2011)

new spines preferentially form towards boutons with pre-existing
synaptic contacts (Toni et al., 1999, 2007; Knott et al., 2006; Nägerl et al., 2007)

B. Intrinsic:

new spines tend to form away from pre-existing spines on the
dendrite (Fu et al. 2012)

dendrites with lower spine densities display a higher degree of
spinogenesis (Holtmaat et al. 2005)

A proposed mechanism extrinsic to the dendrite that can lead to spinogenesis is glutamate spill-over. Conceivable mechanisms intrinsic to the dendrite that can control spinogenesis can be the competition for resources (structural proteins, mRNA etc).
Dendrites of pyramidal cells in some areas of the cortex and the hippocampus exhibit a relatively high turn-over of spine formation and elimination (Holtmaat et al. 2005, Attardo et al. 2015), at least as compared to axonal boutons (e.g. de Paola 2006). Whereas some of these new spines stabilize, many disappear soon after their formation. This indicates that, at least in part, their creation is not fully specified by the existence of a pre-synaptic partner, if we assume that the pre-synaptic partner continues to "attract" them (and that's a big if). The relative independence of spinogenesis from presynaptic activity is further corroborated by the fact that most new spines lack a synapse (Knott et al., 2006).
It appears, then, that dendrites over-produce spines in order to sample their environment for potential synaptic partners. The advantage of such a mechanism can be seen when considering the potential wiring diagram changes that neurons can achieve with such relatively cost-less microscopic changes (Stepanyants et al. 2002). It is unclear at the moment at what point between spinogenesis and synapse formation presynaptic activity becomes an influencing factor in the intact adult brain. A theory incorporating the existing information on spinogenesis is forthcoming.

References

Attardo A, Fitzgerald JE, Schnitzer MJ. (2015) Impermanence of
dendritic spines in live adult CA1 hippocampus. Nature,
523(7562), 592-596. https://doi.org/10.1038/nature14467

De Paola, V., Holtmaat, A., Knott, G., Song, S., Wilbrecht, L.,
Caroni, P., & Svoboda, K. (2006). Cell Type-Specific Structural
Plasticity of Axonal Branches and Boutons in the Adult Neocortex.
Neuron, 49(6), 861–875. https://doi.org/10.1016/j.neuron.2006.02.017

Fu, M., Yu, X., Lu, J., & Zuo, Y. (2012). Repetitive motor learning
induces coordinated formation of clustered dendritic spines in vivo.
Nature, 483(7387), 92–95. https://doi.org/10.1038/nature10844

Holtmaat, A. J. G. D., Trachtenberg, J. T., Wilbrecht, L., Shepherd,
G. M., Zhang, X., Knott, G. W., & Svoboda, K. (2005). Transient and
Persistent Dendritic Spines in the Neocortex In Vivo. Neuron,
45(2), 279–291. https://doi.org/10.1016/j.neuron.2005.01.003

Knott, G. W., Holtmaat, A., Wilbrecht, L., Welker, E., & Svoboda, K.
(2006). Spine growth precedes synapse formation in the adult
neocortex in vivo. Nature Neuroscience, 9(9), 1117–1124.
https://doi.org/10.1038/nn1747

Kwon, H.-B., & Sabatini, B. L. (2011). Glutamate induces de novo
growth of functional spines in developing cortex. Nature,
474(7349), 100–104. https://doi.org/10.1038/nature09986

Nägerl, U. V., Köstinger, G., Anderson, J. C., Martin, K. A. C., &
Bonhoeffer, T. (2007). Protracted synaptogenesis after
activity-dependent spinogenesis in hippocampal neurons. The Journal
of Neuroscience, 27(30), 8149–56.
https://doi.org/10.1523/JNEUROSCI.0511-07.2007

Richards, D. A., Mateos, J. M., Hugel, S., de Paola, V., Caroni, P.,
Gahwiler, B. H., & McKinney, R. A. (2005). Glutamate induces the
rapid formation of spine head protrusions in hippocampal slice
cultures. Proceedings of the National Academy of Sciences,
102(17), 6166–6171. https://doi.org/10.1073/pnas.0501881102

Stepanyants, A., Hof, P. R., & Chklovskii, D. B. (2002). Geometry and
structural plasticity of synaptic connectivity. Neuron, 34(2),
275–88. Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/11970869

Toni, N., Buchs, P. A., Nikonenko, I., Bron, C. R., & Muller, D.
(1999). LTP promotes formation of multiple spine synapses between a
single axon terminal and a dendrite. Nature, 402(6760), 421–5.
https://doi.org/10.1038/46574

Toni, N., Teng, E. M., Bushong, E. a, Aimone, J. B., Zhao, C.,
Consiglio, A., … Gage, F. H. (2007). Synapse formation on neurons
born in the adult hippocampus. Nature Neuroscience, 10(6),
727–734. https://doi.org/10.1038/nn1908

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text:
How should I ship plasmids?
I shipped 10 µL of my vector miniprep to a collaborator in a 1.5 mL eppendorf parafilmed shut and stuffed into a 50 mL conical with some paper-towel padding.  However, something happened on the way and there was nothing (no liquid) in the tube when it arrived.  They didn't make any comments about the microcentrifuge tube popping open or broken parafilm, so nothing crazy happened but something did.
What's the most reliable way to ship plasmids?

Summary

the 10 uL of plasmid miniprep may have been splattered in the cap of the tube (AnnaF)
the eppendorf tube may have depressurized during air shipment and allowed the 10 uL to escape and evaporate
solution: try air-drying or blotting (Jonas) your minipreps prior to air shipment

Details
As AnnaF wrote, the 10 uL of your plasmid could have been hidden in the cap or dispersed around the tube, making appear empty.  You should check with your collaborators to be sure they centrifuged it.
According to a fedex document on shipping perishables (pdf) and a paper measuring the temp and pressure of air shipments (pdf), fedex and ups air shipments may experience low pressure environments around 0.56 - 0.74 atmospheres (atm).  At these relatively low pressures, perhaps an eppendorf tube sealed at 1 atm might breach.  The papers also note that ground shipments that pass over the rockies (i.e. in Colorado) may experience ~ 0.64 atm.
So perhaps your 1.5 ml eppendorf tube depressurized during the shipment?
It would be interesting to do some tests on the pressure-worthiness of eppendorf tubes.
Regarding the original question, in 2007 I prepared and shipped (fedex) a library of thousands of minipreps to hundreds of users.  1uL of miniprep was dispensed into wells of 384-well plates and airdried, then sealed with aluminum, then mailed.  Users rehydrate a well with 10 uL of water.  Generally it works.

A quite safe way of shipping plasmids is to put them on filter paper (see protocol and send a letter. Much cheaper.

+1 Fast, easy and very stable even for overseas shipments.

Plus, customs are much less fussy about a piece of paper than they are about anything liquid.

This is the best way to go. You can also wrap the filter paper in a bit of cling wrap just in case.

Did they try to centrifuge the tube when it got there to push all the liquid to the bottom? I know that especially when working with such little amounts that even shaking it up a little can disperse the contents all over the tube.
We have received plasmids from other labs before. Generally speaking the plasmids are sent in Screw-cap microcentrifuge tubes inside of some sort of canister. This is then packaged in a little foam cooler with dry ice to keep everything cold.

Since tubes can be crushed in the mail, the safest way to ship plasmids is to drip a few uL into a filter paper, and then wrap it up to seal it with parafilm, and fill out a detailed form about its content.
Good luck.

We have gotten into similar situations when other labs have sent us plasmids (or when we have taken out ancient tubes out from storage boxes at -20), and have since adopted the filter paper method. Another point to note is that you can just add, say, 10 μL of water to the "empty" tube (Mac) and use 1 μL for a transformation. It has always worked for us. DNA is seemingly invisible!

I will also strongly recommend blotting a microgram of your plasmid DNA on a piece of filter paper (the filter paper is important for extraction on the receiving end). Also, it is very very helpful if you clearly indicate the amount of DNA (approximately) and circle the blotted DNA in pen so there is no ambiguity about cutting out the circle of filter paper containing the DNA.
On the receiving end, you should always freeze the paper right away or elute the DNA off of the paper in TE buffer or water and then freeze. Failure to do so will eventually lead to DNAase degradation of the plasmid DNA and then you'll have to wait for another shipment.


text:
What is the reason behind choosing the reporter gene when experimenting on your gene of interest?
I noticed within example experiments in class that different reporter genes are chosen to be inserted near your gene of interest to prove whether or not the gene is being expressed. For example, you may insert the gene for fluorescence next to your gene of interest so you know if it is transcribed or not by whether the organism's cells are fluorescent and to what degree they are fluorescing at.
I have noticed in some experiments that have multiple versions that in one case they use the fluorescent gene and in the next a different gene (for example lactose). Both portions of the experiment use almost the exact same steps so why would they not choose the same reporter gene?

Already good answers here. Btw, I'm keeping a blog about reporter genes if you are interested.

Different genes will serve different purposes. For example, if you want to perform colocalization studies, then fluorescent genes like eGFP and DS-Red (or any variation of those, namely Emerald, mCherry, etc) will be quite useful, since you can use different filters on your microscope for the various fluorophores. For morphological assessments, perhaps a LacZ gene or alkaline phosphatase may be better suited, since the detection of these enzymes is compatible with standard histological techniques and you get usually much better contrast than with fluorescence. For quantification purposes, you would use probably firefly luciferase or CAT, since there are very good quantitative assays for these enzymes.
The decision will also depend on the degree of experience of the lab with each detection technique and even in the availability of the constructs. If a reporter is already made by the lab next door and if it fits your needs, you'll just use it, instead of creating your own, even if the reporter gene would not have been your first choice.

Reporter genes are also used when the experiment is trying to distinguish the effect of that gene that the scientist introduces (exogenous) and the same gene that the cell already has (endogenous). Ideally, one would like to choose a cell system that does not contain endogenous genes of interest because of this confounding effect, but it is not always feasible.
Reporter genes are also mainly used when it is impossible or very difficult to directly measure a gene of interest, especially if the gene of relatively obscure in the scientific literature. For example, cells that do not express a thymidine kinase gene are impossible to visualize by Positron Emission Tomography. It's not because there is anything wrong with the gene of interest, just that the cells need a way to retain a radioactive agent that can be detected by the PET machine. Likewise, fluorescent genes are very often used because there is often a need to see where the gene product is located within the cell, and the only way to do that is to combine it with something visible by microscopy which lets you see fluorescence and the cell structures. Lastly, a reporter gene is often used when the gene of interest is hard to detect because of limitations of commercial antibodies or detection kits, and the reporter, being very commonly used, is easy to detect with commercial products and antibodies.

Reporter genes have become an invaluable tool in studies of gene expression.A gene consists of two functional parts: One is a DNA-sequence that gives the information about the protein that is produced (coding region). The other part is a specific DNA-sequence linked to the coding region; it regulates the transcription of the gene (promoter). The promoter is either activating or suppressing the expression of the gene.
The purpose of the reporter gene assay is to measure the regulatory potential of an unknown DNA-sequence.So it is quite helpful,it;s kind of reverse engeneering the function and then let know what exactly it does.

This answer addresses what a reporter gene is, but it seems like the OP already knows that. The OP is actually asking why you would use a particular reporter gene in a particular context. If you have additional insight about this question that has not already been included in the two posts above, please revise your answer.


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How many times did endosymbiosis occur?
According to the endosymbiont theory, mitochondria and chloroplasts originated as bacteria which were engulfed by larger cells. How many times is it estimated that this occurred in the past? Are there any examples of this process being observed directly?

What do you mean "how many"? Do you mean how many time it should have happened before mitochondria became our endosymbionts? This question doesn't make any sense for me...

I think this is a good question. Was it a unique or rare event, or an inevitable consequence of bacterial colonies cooperating? This is a fundamental issue in the development of multicellular life.

Well, it seems quite obvious that it was not a single I-eat-you-but-you-survived act but rather a convergence of endosymbiotic and host species into a greater and greater cooperation.
Of course this leaves a question if there was one or more species of endosymbionts involved.
Mitochondria are a very primeval story forced by the oxygen catastrophe, so it is hard to say, although great majority of mitochondria seems to have a single origin.
Plastids are much more divergent, however it seems that they did originated from a single source, diverged into chloroplasts, cyanelles and rhodoplasts and were later mixed up by numerous acts of secondary and even tertiary endosymbiosis (plus a further evolution); this variety can be especially seen within Euglenas, and they are the main group investigated in this manner.

what is the story of oxygen catastrophe which put some force upon mitochondria?

Long and speculative -- in short, aerobic bacteria are postulated to be ingested by larger, anaerobic ones which required something to reduce oxygen level inside its cells. I think you should ask a new question about it.

@Vass:  you might look at http://www.amazon.com/Oxygen-Molecule-World-Popular-Science/dp/0198607830/ref=sr_1_4?s=books&ie=UTF8&qid=1336399813&sr=1-4 It is fairly tough going for a popular science book, but is about how life learned to live with and use oxygen

I think @mbq has covered the frequency question better than I can.
There is at least one modern example of this kind of new organelle formation.  Aphids have a deep, intracellular endosymbiont Buchnera involving some genome transfer that has developed in the last 200 million years.
There are many articles about this topic (eg: Nature from 2000), and it was a little controversial 15 years ago.  Now, it's largely accepted that this is a modern development of endosymbiosis, and a confirmation of the theory.

@Thomas Ingalis, with the example of Buchnera, why does a intracellular endosymbiont involve genome transfer? What is that referring to exactly?

@Vass, to be correct, I should say functional redundancy. Nikoh et al published an analysis in 2010 screening a complete aphid genome for transfered genes.  From the abstract: "Our results excluded the hypothesis that genome reduction in Buchnera has been accompanied by gene transfer to the host nuclear genome, but suggest that aphids utilize a set of duplicated genes acquired from other bacteria in the context of the Buchnera–aphid mutualism."  This redundancy can be construed as a step to organelle formation.

It depends of what you call endosymbiosis. In the sense of mutualistic interaction between host cell and intracellular organism, it also include Rhizobium bacteria and Fabaceae plants, some Cnidaria and algea in their cells, and even some micorrhizal fungi that invade into plants cells. But parasitic interactions are also sometimes call symbiosis, as symbiosis means just “living together” and the balance between profits and costs for each partner are sometimes difficult to measure and even changes in time. So in this wider meaning even Trichinella worm that live in mammal cell is an endosymbiont and endosymbiosis is very common.

There are evidences of secondary endosymbiosis i.e. organelle within an organelle. This is quite evident in Chromaveolates. Many unicellular Chromaveolates which had been classically referred to as unicellular Algae, have chloroplasts derived from other algae. This organelle in turn has a membrane bound suborganelle. For a quick reference you may see this article. Euglena and diatoms are some famous examples of this phenomenon.


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How to reduce edge effects in cell-based high-throughput experiments?
In high-throughput experiments where cells are cultured, treated, stained, and imaged in 384-well microplates, I frequently see significant edge effects. For instance, the following plot shows cell count (as measured by CellProfiler from images) for each well on such a plate:

How can I reduce these effects? I can try to correct for them statistically, but it would be so much nicer to reduce them at the wet-work stage.

Lundholt et al. describe a very simple trick: let the plates settle in air at room temperature for a couple of hours before placing it in the incubator. We tried it out, and although the color scales are different in the following plots, it is easy to see that it really does help:

(Thanks to Sigrun Gustafsdottir and Kate Madden for testing this.)
I have seen some people "pad" the plate by using only the 308 interior wells and filling the  76 edge wells with only liquid, but I have no data for how well that works.
Allison Tanner from Corning has collected a compendium of techniques to reduce the occurrence of irregular patterns of cell adhesion or “edge effect” in microplates. Brifly summarized:

Avoid bubble formation in the media when seeding cells.
Use cell harvesting methods that foster adequate mixing of the cells in the media and complete dissociation of the monolayer into single cells.
Seed cells at a density that will support attachment and growth.
Allow cell attachment to occur in media of the proper formulation for the particular cell type.
Dispense cells in an appropriate volume of media.
Follow strict aseptic techniques according to respected microbiological practices.
Cultures should be routinely monitored for mycoplasmal contamination.
Minimize entrance to incubators to help reduce fluctuations in the internal environment.
Allow cells to settle in the microplates while the microplates are on the benchtop.
Culture cells in an environment that reduces the formation of static electrical charges.


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Has anyone tried Gibson Assembly Optimizations?
Has anyone tried the chew back-anneal in vitro DNA assembly method (known as Gibson Assembly) for difficult sequences, like GC-rich sequences? How big constructs could you efficiently assemble? Did you use the original protocol or some optimizations?

I've just joined the Gibthon team, so if you need any more specific help let me know.

I haven't tried Gibson Assembly myself, but I do know the Cambridge iGEM Team in 2010 and 2011 developed a web-based tool - Gibthon - for automatically generating primers for Gibson Assembly.
You could try running your difficult sequences through their tool to see what it suggests for primers.  It apparently gives you a "value for the Gibbs free energy of the worst case secondary structure of the primer."

Thanks. Yes, I have heard of the Gibthon software, haven't tried it yet. We have been using the IDT predictor of secondary structure of the primers. Might be worth comparing it to the Gibthon tool. It would have been nicer if some optimizations of the reaction conditions make the use of such predictors obsolete. Not to mention that their precision is questionable.

I also haven't tried it, but theoretically GC-rich sequences will interfere with Gibson assembly. Since unique homology between the ends is key to the assembly process, repetitive sequences which cause non-unique overhangs will increase the possibility of incorrectly ordered assembly. One solution would be to use alternative codons where possible to decrease repetitiveness.
I'm sure all the overlap design tools will try to avoid repetitive overlaps. Prof Jim Hasselhoff at Cambridge Plant Science is a big proponent of Gibson assembly, it was one of his students who wrote Gibthon. If you email him I'm sure he'll give you specific advice.


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What is the optimal frame size for protein secondary structure prediction methods?
The whole question is

What is the optimal frame size for the second and third generation protein secondary structure prediction methods? Justify your answer.

I remember it has something to do with the average length of alpha-helix. More specifically, 3 on both side of a site. So in total the frame length should be 7. But I can't remember the reason behind the argument.
What do you think?

According to what my professor said in class, 2nd and 3rd generation of protein secondary structure reconstruction relies on statistics data of several consecutive residues.  I guess what he meant by "frame size", is how many adjacent residues we should take into account in the algorithm.

By frame size, do you mean sliding window?
I know that if you want to predict a secondary structure of a transmembrane protein, then your window size should be 20 amino acids (this is the average length of 1 transmembrane alpha helix spanning through the membrane).
I found this paper by Chen, Kurgan, and Ruan[1].
It basically says that the window size depends on what kind of pattern you are looking for, but in general, 19 residues should be optimal.
Also, secondary structure predictors rely on many features like hydrophobicity, missing coordinates in X-ray structures, B-factors, motifs, etc.

Chen K, Kurgan L, Ruan J. 2006. Optimization of the Sliding Window Size for Protein Structure Prediction. CIBCB '06: 2006 IEEE Symposium on Computational Intelligence and Bioinformatics and Computational Biology, pp.1-7, 28-29, doi:10.1109/CIBCB.2006.330959.

That's the problem of computational biology - sometimes it's difficult to find what exactly what a term means.  But I think my professor asked for general protein secondary structure predication, not transmemebrane protein.

OK, so 2 responses, each concerning different protein segments of different types/functions/structures, but not an answer really getting to the heart of the question. Because of the homework tag, I am tempted not to reply - this is something the student should accomplish on his/her own. So, I'll give a general answer to start you thinking. Basically, such a value - be it 7 or 20 for TMs or 13 - is determined empirically.

I found that 13 worked best for Neural Nets and SVM performance of secondary structure information when I ran this in R.

Any reason behind it?


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What are the main mechanisms of interaction between the nervous and immune systems?
We know from pop science that our psychological states have an effect on our immune systems ("worrying ourselves sick", etc.), but what are the actual mechanisms through which our nervous systems pass information to the immune system?
Cell mediators come to mind, but where in the body would a nerve cell release an interleukin or other factor? (Put another way, are neurons releasing these factors as a part of their normal cell metabolism and the side effect is a communicative effect with the immune system?)

It is not only the immune system that prevents us from getting ill. Worrying much in my opinion won't make you catch a cold; rather, you can get problems with your cardiovascular system (arrhythmias, hypertension, angina pectoris) or limbic system (panic attacks, sweating attacks etc.).
The connections are many; here are some possible ones:

There are many glands in our body that are either densely innervated (like the thyroid gland) or are a part of nervous system (like pituitary gland, epiphysis). They can release hormones directly to our blood and make our hormone profile unstable, hence also impacting the immune system.
Sympathetic/parasympathetic nervous system is under direct control of the CNS (N.vagus, carotide sinus sinus etc.) and can dramatically change the peripheral resistance of our vascular system by constricting (=> angina pectoris, hypertension) or dilating (swellings, bowel movements problems -> constipations etc.) the small vessel. This alone can start so many different mechanisms resulting not only in being ill, but also being dead.

Speaking about connection between the nervous and immune system we must consider that:

Many neuromediators, like adrenaline, for example, have clear effects on immune cells.
The mediators and humoral factors released by nerve endings can easily diffuse outside of the synaptic cleft and act locally, attenuating immune response.

Here's pretty much the first thing that popped up on pubmed:http://www.ncbi.nlm.nih.gov/pmc/articles/PMC3144148/
 So clearly there are direct neurotransmitter receptors on immune cells themselves. Pick a favorite neurotransmitters and search "xyz and the immune system".
However, I can't tell you how relevant that is in terms of actually affecting large scale shifts in immune function. I think the previous poster Alexander Galkin had it right in that the nervous system affects hormones which have the ability to coordinate large scale, complex immune system shifts. It's like the president calling the generals to organize some troop movement and how many troops to deploy, what their targets are, etc.
I just wanted to add that tidbit that our "soldiers" do in fact carry cell phones. If the president wanted to, he could call the troops personally.

Yeah, I definitely realize there's quite a bit of info out there about it. :)   This question was from the early days of the site when we were trying to get a variety of content out there.  Great addition to the other answer!


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What are the mechanism binding histone code with alternative splicing?
There are some bioinformatical considerations showing that there is a significant correlation of some between histone code and alternative splicing; but what are the hypothesis about the underlying mechanisms? Is there any experimental insight in the problem?

According to literature, histone marks seem to be affecting splicing both directly and indirectly.
The indirect regulation appears to be due to biophysical hindrance, where a certain chromatin structure would cause Pol-II slowdown, allowing the splicing machinery to do its job.
The direct regulation seems to be essentially dependent on H3K36me3 and H3K4me1, marks that are recognized by MRG15, which in turn recruits the splicing regulator polypyrimidine tract-binding protein to splice sites.
Ref: Luco, R. F. et al. Regulation of alternative splicing by histone modifications. Science 4 Feb 2010


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What are the limiting factors for gene length and number of exons?
I recently downloaded gene annotations for Homo sapiens from Ensembl for some bioinformatic analysis. The vast majority of the gene annotations have 20 exons or less, although there are some that have as many as 250. I know enough about gene annotation to take these predictions with a grain of salt, but it got me thinking...what are the biologically relevant factors that might limit the length, exon number, etc of a gene? Is there any real possibility for a gene to have 50 exons? 100 exons? 250 exons? From a biological standpoint, where is the line drawn and why?

How did you retrieve the genes/exon annotations? BioMart, or EnsEMBL API?

I got the compressed (gzip) GTF files directly from the FTP site with wget.

The EnsEMBL Perl API is a cool way to retrieve the precise data you need - http://www.ensembl.org/info/docs/api/index.html

This question drops firmly into the lap of molecular evolution and the constraints that are placed upon genes by the forces of mutation, selection, drift and recombination.
There are numerous situations, particularly gene duplication, that can result in a gene that is free from the selective constraints of it's parent, many of which will accumulate so many deleterious mutations as a result of stochastic processes that they will become non-functional e.g. psuedogenes. Some can be altered and rearranged, accumulating exons and introns, and if they infer a fitness benefit on the organism, may be moved to fixation within a population.
Evolution is a population genetics process, and there are many variables which can effect the outcome, not least the difference in populations size. The genomes of larger populations (such as those of bacteria) appear to have much smaller genomes, and of course no (at least not spliceosomal) introns, perhaps as a result of increased fitness due to the decreased generation time of an organism with a more slender genome. It would be a good idea to read The Origins of Genome Architecture by Michael Lynch, as I think he answers your questions, better than I can.
Many of the genes you retrieve from EnsEMBL will of course have experimental evidence to support them. The genes that are predicted in the pipeline can be looked upon with less confidence, but you can of course look at the alignments with closely related species to see if you think the introns/exons are indeed viable. An example of a gene with 79 exons is the Dystrophin (DMD) gene, the longest annotated gene at 2,217,347bp (see Roberts et al, 1993 and Nishio et al, 1994).

I doubt there is really any direct limitation; the best test would be to check if the size matches, i.e. that 1kbp gene with 100 exons would rather have to have way too short introns.
Quick search over NCBI Genes show even a 317-exon gene, although all those edge cases seem to be some unclear siblings of titin which is just huge on its own.

Talking about annotation quality control, one needs to think about the molecular constraints on the size of an exon, or intron for that matter. The (theoretical) minimum length of an exon would have to 1bp, although one would also need to think about the binding of molecular machinery involved in exon-intron boundry recognition and the splicing of adjacent exons. I should think exons of less than 6bp would probably not be considered functional? See http://www.jbc.org/content/270/6/2411.full and http://mbe.oxfordjournals.org/content/23/12/2392.full

Agree with mbq - titan is the longest gene I know of and it has well over 100 exons. Titin and dystrophin are well characterized genetically and not predictions.  titin is the champion exoner with 363 exons.
Its only examples like this that can allow the gene predictors to run on as long as they do I think as the predictions are trimmed heuristically to resemble the known gene structures/lengths/junctions etc.

Dystrophin is the longest gene, but doesn't encode the longest product, which of course is Titin. 363 exons is one hell of number! Lol :)


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How to understand influenza strain designations?
What do the strain designations for flu mean?
For example avian flu is classified as H5N1, what do the letters H, N and numbers 5, 1 mean? Is it more than a simple string-identifier?

The sub-type is named for the broad classes of the hemagglutinin (HA) or neuraminidase (NA) surface proteins sticking through the viral envelope. There are 16 HA sub-types (designated H1 - H16) and 9 NA sub-types (designated N1 - N9). All of the possible combinations of these influenza A subtypes infect birds, but only those containing the H1, H2, H3, H5, H7 and H9 and the N1, N2 and N7 surface proteins infect humans, and of these, so far, only H1, H2, H3 and N1 and N2 do so to any extent.
Read more: http://www.fluwiki.info/pmwiki.php?n=Science.NamingInfluenzaViruses

Subtype is denoted using the HxNy notation for the variant of hemagglutinin and neuraminidase.
Strain names are specified as: [virus type]/[location of origin]/[sequential number of isolation]/[year of isolation] ([subtype]), such as "A/Alabama/AF2070/2010(H1N1)"

Virus type would be A, B, or C for the various forms of influenza (influenza A being the most common among humans)
Location can be a country, state or city (e.g. Laos, Alabama, or Moscow)
The number of isolation can be a simple number, but sometimes it goes beyond; e.g. "AF2060" or "UR06-0455", possibly due to the tremendous number sequenced nowadays.
Year, obvious.
The subtype refers to the aforementioned HA/NA subtype.

Many more strains can be found on the Influenza Research Database.

The two letters represent the type of hemagglutinin (H) and neuraminidase (N) present on the viral surface. Those are the major surface proteins of the influenza virus and therefore crucial for the immune response.
From Immunology and Evolution of Infectious Disease

HA and NA reactivities with antibodies define the subtypes of
  influenza A (Cox and Subbarao 2000). Fifteen different HA antigenic
  subtypes occur, each subtype cross-reacting relatively little with the
  other subtypes. Nine distinct NA subtypes are known. The designation
  HxNy describes an influenza subtype with HA antigenic subtype x =
  1,...,15 and NA antigenic subtype y = 1,...,9.

The numbers represent different subtypes of the hemagglutinin and neuraminidase proteins.

Adding to the other answers posted:
Hemagglutinin  (HA or H) -  Helps the virus enter the cell by binding to sialic acid receptors on cell membrane. It then unfolds in a lysosome and fuses the viral and lysosome membranes.
Neuraminidase (NA or N) - Helps the virus exit the cell by cleaving the terminal sialic acid residues at the progeny virus release.


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Transmission of epigenetic regulation through surrogate mother
I know the basics of epigenetics, but I do not know how epigenetic mechanisms are transmitted from parents to children (or if there is even enough literature to derive a consensus). My question is relates to surrogate motherhood, in which an egg is fertilized in vitro and then implanted in the womb of a woman other than the egg donor.
Are all the factors that affect epigenetic mechanisms encoded in the genome? Is it possible for the surrogate mother's epigenetics to influence the epigenetics of the child/children?

Epigenetic information is information that can be inherited through cell division that is not encoded in the DNA sequence. This includes, but is not limited to, DNA methylation and histone modifications (there is also non-chromatin based epigenetic information). A nice example is the centromere, the chromosomal region that binds the kinetochore and is important to attach chromosomes to the mitotic spindle during cell division. The location of the centromere on the chromosome is encoded by a specific nucleosome composition and does not seem to rely on the DNA that wraps around those nucleosomes: The DNA sequence at centromeres is not even conserved from chromosome to chromosome (with the exception of budding yeast), and there are several known examples of people and families where the centromere is in a different place. However, all the mechanisms for maintaining this epigenetic mark are encoded genetically.
As epigenetic information is transmitted through cell division, it is directly inherited from the biological mother (with the exception of e.g. the centromere, most epigenetic marks from the father's chromosomes are removed when sperm is created).
However, (some) epigenetic information can be modified. This is obviously important during development where gene expression patterns of a liver cell need to be stable but different from gene expression patterns of a neuron, even though both descend from the same cell.
There is evidence that the metabolism of the mother will influence the epigenetic program of the child; diet being one of the determinants. It has also been suggested that epigenetic changes may influence behavior. Thus, it is indeed possible for the surrogate mother's epigenetic state to influence the epigenetic information of the children.

The idea of epigentics is that there is more info in chromatin than just in the DNA sequence it contains and that this info can be altered in vivo.
Thus, one cannot inherit epigenetic changes from a surrogate (which brings no new physical genetic material), but the fetus may develop some due to its interactions with her.

The definition of an epigenetic mark I work with is the following:

impacts gene regulation without touching the DNA sequence,
is reversible,
is inheritable.

With this definition in mind, you could say that only DNA methylation is a true epigenetic mark, leaving histone modifications and the like to canonical transcription regulation machinery. Yet it really depends on what scale you place yourself. Epigenetics at the scale of the cell is not the same thing as at the scale of the organism.
So if a surrogate mother can alter the epigenetic state of the pup or child or foetus, it will not be due to transmission of information in the traditional sense, but rather be due to her physiological state in the case of the foetus (many hormones act through nuclear receptors that can recruit chromatin remodelers), or as mbq mentioned it, due to her behaviour in the case of the child or pup.

X inactivation is not due to DNA methylation but does impact gene regulation without touching the DNA sequence. Also, the inheritance of the location of the centromere would not fall under your definition, so what kind of mark would that be?

You are totally right, for some reason histone variants did not come to my mind when answering :)


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What is a good miniprep protocol for the class room?
I'm trying to find a good protocol for plasmid minipreps and I'm looking at 3 preps I've found:

Using phenol/chloroform

extract with phenol:chloroform:isoamylalcohol,
isopropanol precipitation, 12,000g spin down,
rinse with cold 70% ethanol.

Using lysozyme

lyse with lysozyme,
remove pellet,
isopropanol precipitation,
wash with cold 80% ethanol.

Using alkaline lysis

open cells with 80% glucose in EDTA buffer,
add SDS and NaOH,
pellet protein/membrane with acetic acid/acetate,
ispropanol precipitation,
wash with cold 70% ethanol.

They all differ in how to break open the cells and separate plasmids from the rest of the cell -- quite a bit.  Can anyone help me figure out which protocol is best here?

Are you sure you want to have students use phenol? Most labs probably use kits for DNA extraction (e.g. Quiagen MiniPrep). It's much easier and also safer.

i'm going to mark Reven's answer as the best since it helps me most choose amongst the protocols.  but thanks for pointing out how cheap the qiagen kits are- i'd thought they were much worse.  you do learn something by using a protocol, but dang the columns are easy!

FYI while its true about $1-2 per pre for spin columns, but the reagent cost for SDS/NaOH is about 40 cents per prep. you have to do over 100 preps to see the savings though.

Just wanted to update that i bought columns from biobasic and while it is not as for instruction as these basic protocols, they are certainly cheap enough and should be considered for the classroom - upvotes all around.

@MadScientist Spin columns are indeed the most common option, but there is a pedagogical angle: There isn't much to learn about following a kit manual, but a PCI extraction teaches some important biochemistry. Even if routine purifications are done with spin columns, PCI can be a valuable fallback when you just so happen to get stuck because some impurity keeps getting through your column.

in the end i talked about what each step of the spin prep does and it works fine.  the old methods are not really much more rigorous when you talk about pH changing affinity on silica, etc

In my experience, the P:C:I method will get you higher yields, and is a bit more simple (in steps and chems involved), but as @Mad Scientist has said, phenol use might be an issue. It depends on the age of your students.

It is not only the age of the students, it also depends on the available facilities. You DO need a hood for instance, and you need to have a service available for the disposal of phenol.

Phenol and chloroform are highly toxic and carcinogenic and should definitely not be used by students or outside of a proper fume hood.

The protocol I used in my genetics lab course was alkaline lysis followed by ethanol precipitation similar to this one.  Nothing terribly toxic or requiring a fume hood (at least at the small volumes used).
The more practical, reliable method is to use a miniprep kit (usually spin columns) from one of several suppliers (Qiagen, Promega, Bio-Rad, etc.) which are fairly affordable at $1-2 per prep.

If I recall correctly, alkaline lysis is the underlying method behind Qiagen DNA purification kits (miniprep, etc). With the spin columns from the kit, you get to cut out the alcohol precipitation and drying. Maybe doesn't sound like much, but it is worth the price of the kit because it eliminates more than 10 minutes and a sensitive step from the protocol. My two cents: you do not want to use phenol in the classroom.

All the prep kits I've used (Qiagen, Promega, Invitrogen) do use alkaline lysis.  I like the columns just for the sake of not having to worry about totally evaporating all the ethanol (I use them with a vacuum manifold so I suck for an extra minute or two to let it all evaporate) or looking for some half-invisible pellet.

Liking this more once i found the qiagen recipes on Open Wet Ware.. http://openwetware.org/wiki/Miniprep/Qiagen_kit makes me feel better about using it in a class.

I think the spin column kits are the way to go. As mentioned already, the benefits of the kits are that they are easy, safe, and (most importantly) how almost every actual lab does plasmid purification these days.
The biggest criticism of the commercial kits is that you can get by easily without knowing anything about what is actually happening in the tube. However, as noted in the comments to Nick T's answer, there is no secret proprietary technology in the kits--they use the alkaline lysis method. This means that you can still teach the mechanism in the class while getting the benefit of the spin columns.
One option, for teaching purposes, is to make all of your own solutions and just use the spin columns on the supernatant after pelleting the precipitated membrane/protein. That way, you get the benefit of insisting on real names for the solutions (NaOH is a more educational name than P2) and you get to use the spin columns in place of the ethanol precipitation. I believe there are cheap sources that just sell the spin columns, although I have never used them. You want to avoid the ethanol precipitation because it adds a bare minimum of 30 minutes to the protocol (for my protocols, it was more like an 1.5 hours or more). And waiting for ethanol to evaporate so you can resuspend is like watching paint dry unless you have a speed vac.
Yield shouldn't be a big issue in a class situation, but you can boost yield with the column by prolonging the final elution step. I think the given instruction is to let the TE buffer (Tris) "elute" for 1 minute before the final spin, but I have routinely let the TE sit for about 15-30 minutes before spinning down. The difference in yield blew out the exposure on the gel camera.
There are certain cases where phenol:chloroform extractions are still desirable, but this is definitely not one of them. I used to do P:Cs routinely when working with RNA and yeast genomic DNA. I have never seen anyone do anything other than a miniprep when preparing plasmids. What's more, the hassle of working in a fume hood and the risk of phenol burns encouraged a lot of people in the lab to prefer the spin kits even when working with genomic DNA.

after some effort we did move to the spin kits...

Using spin columns, my professor and I extracted ~30 plasmids from ~30 samples in a few hours.  It's easy, cheap, and it yields good quality plasmid.  My sample was quantified by UV spec and it was at about 200ng/ul.

To save cost, you can buy spin columns alone from suppliers such as Syd Labs (http://www.sydlabs.com/spin-column-for-plasmid-miniprep-p110.htm) and make home-made buffers. The suppliers normally provide the buffer recipe.


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What specific membrane adaptations do cells have for saline-rich environs?
Are the cells of marine animals and flora equipped with special ion exchange pumps to mitigate the effects of a saline-rich environment?
Or have the cell's membranes adapted through structural changes to counter the osmotic forces?

This 1969 Steensland paper seems to suggest that the membranes of halophiles are stabilized by sodium ions and they rapidly denature at lower-salt conditions (2.2 vs. 4.3 M).  The protein composition of the membrane was generally acidic, stabilized by all the Na+.
As far as what the role of the halophile membrane is in sheltering the cell from the high ionic strength solution seems left unsaid.


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Bacterial chromatin binding data?
I'm looking for data - maybe CHP^2 data that shows chromatin binding to a prokaryotic genome under some specific conditions.  Can anyone point me to a source?

Well, a byproduct of the fact that prokaryotes do not have nucleosomes, is that they don't have chromatin either. Compaction is achieved through supercoiling of the genophore.
Do you mean binding of a protein to prokaryotic DNA ?

@agrimaldi: That is not exactly the case.  Some prokaryotes have nucleosome-like structures.

A large number of prokaryotes do indeed have nucleosome-like structures.  The most well studied is H-NS in E. coli, Salmonella and some other deltaproteobacteria.  H-NS like molecules have also been found in mycoplasma (Lsr2).  One of its roles is to bind AT-rich DNA and silence transcription.  The binding is usually to suppress the expression of foreign DNA which tends to have a lower GC content than the host organism.  There is a great paper that did ChIP on ChIP with H-NS in Salmonella by Navarre et al in Science (Pubmed ID: 1676311).
As a side note H-NS does compact DNA and there is a paper by Dame et al. that demonstrates it with atomic force microscopy.

@agrimaldi: There's a lot of papers about H-NS and H-NS like proteins, it's very interesting.

I was just reading this paper:  In making the case for chromatin binding, to the bacterial genome, it seems like the nucleate at supercoiling regions but then may migrate.  The GC content idea sounds intriguing, but wondering if there is any more specificity the chromatin exhibits upon binding.   Genome biology, 5(12), 252. doi:10.1186/gb-2004-5-12-252


text:
What limits the maximum spacing of Nodes of Ranvier and which organisms tend to have the widest gaps?
Assuming that a longer distance between gaps in the myelin sheath is beneficial for an organism due to the increased propagation speed, what is the limiting factor in determining the maximum spacing between nodes?  I have previously encountered the analogy of the Nodes of Ranvier as forming a circuit between themselves and the previous node in order to pass on the action potential, which leads me into thinking that it is simply electrical resistance that is the cause however I assume the analogy is an over simplification and would be very interested in any alternatives.
Also, is there a significant variation in node separation between organisms?  Which organisms have the greatest gaps?

This is best answered via something called cable theory.  Basically, as the action potential (AP) propagates along the membrane of the axon, it's tripping voltage gated channels that "renew" the flux of ions into the stream at the nodes. There are no ion channels under the sheathing (or they are there anatomically and inactive, but I don't remember), so the current is able to zip through that area.  There's no renewal of AP, but there's no loss due to leak of ions either.  Since there's no influx of ions from outside of the membrane until the next node, the capacitance and resistance of the membrane is exponentially decaying the voltage.  So, if the distance between the nodes was too long, the voltage would just die off and the AP wouldn't propagate.
I don't have any anatomical data about individual species, but know that only the smaller diameter axons require sheathing to compensate for the slower ion flow due to higher resistance (e.g., the squid giant axon(not to be confused with the "giant squid" axon) has no myelination due to its large diameter).

Right on. The only thing I would add is to explicitly mention the distinction between axial resistance and membrane resistance in cable theory. The axial, or longitudinal, resistance is what is relevant here as it determines how much current flows down the center of the axon. As you say, this parameter is mostly affected by the axon diameter (larger axons have a lower axial resistance). Thus, the decay of voltage along a large diameter axon will be less severe than along a small diameter axon.


text:
bi-directional transcription experiment
We suspect a bi-directional transcription event is happening at a locus in our organism where two genes are directly adjacent to each other. The annotation data is not well established. The intergenic distance is probably less than 200 base pairs.
The two genes are expressed in opposite directions towards each other. Base on the preliminary transcriptomics data, it seems like one gene is over transcribing (3' UTR perhaps?) into the adjacent gene, possibly resulting in some kind of transcriptional regulation of the adjacent gene.
Here is a rough diagram of what we think might be happening:
------------------------==========gene A================>----------------------

----------------------------------------<====gene B=====-----------------------

Of course we need to first confirm this by designing primers to see if this over transcription is actually happening.
If this is happening, we intend to do some knock down experiments. We have no transgenesis available in our organism, only RNAi by dsRNA. It is possible to specifically knock down geneA by introducing dsRNA to the 5' region of geneA that does not overlap with geneB. Perhaps this will lead to ectopic/over expression of geneB.
Is there anyway to knock down geneB specifically without knocking down geneA? It looks like designing dsRNA for geneB would knock down both A and B.

What organism is it? And how big the genes A and B are?

Both geneA and geneB have multiple exons. Transcript length-wise, both are around 2kb. Both have long introns resulting in a 4-5 kb total gene length.

Technical issue -- now, your diagram may show almost any possibility depending on the font selected in browser. Use code input (either prefix with 4 spaces or select and use {} button to get monospaced font which will behave consistently).

How well is gene A annotated? Do you have the gene A sequence (after post-transcriptional modifications)? If you do, you can order it from a company like DNA2.0 and they will synthesize it for you for like $0.35/base. Then you can transform the cells with this plasmid and do a knockout of gene B by inserting some sequence at the ~200bases overlap.
Also, I guess you can transform the immediate transcript (without modifications). It will also be modified once it is in the cell. But it will be longer and more expensive to synthesize, but there are a few methods you can use to do that.
Let me know if this idea appeals to you. I can give you more info if you are interested.

If gene B makes a protein product, you can try designing a morpholino against the 5'-UTR of gene B. This can prevent translation initiation at the ribosome as the morpholino occludes mRNA entry into the ribosome. You can detect this by western blotting.
If gene B makes some sort of regulator RNA, you will need to target expression of gene B at the DNA level. If gene B has well established splice sites, a morpholino can interfere with splicing enhancers and silencers, and therefore, should produce a non-functional or impaired RNA product. You can detect splice variants by amplifying gene B from cDNA.
Theoretically, the advantage of using a morpholino is due to it's single stranded nature. Morpholinos designed against gene B should have no effect on gene A due to non–complemenetarity.


text:
Do human beings have pheromone receptors?
What is the current consensus on whether or not humans have receptors that detect pheromones?
If there are purported receptors, in what anatomical areas are they located?  With what organ systems do they interact?

It would be the vomeronasal organ... I'm no expert in human brain anatomy, but I think there is a bit of a controversy on whether a functional one exists in humans...

I think it is perfume industry that knows the ultimate answer. Although some smells are sexy and attractive, there is no human pheromone as far as I remember reading (and smelling).

From reading popular science articles on the matter, it appears to be vestigal (measurable in careful studies, but swamped by other factors when looking at how it affects the relevant behaviors)

It's my personal hypothesis that pheromone receptors (wherever else they exist) also exist on the sides of the part of the nose directly between the nostrils, as well as in the ears. But I have no proof of that for you!

From what I have gathered, I would think that humans do have receptors that are able to detect pheromones.
For example, some studies that have indicated human responses to pheromones.  Of these pheromone responses, some have been traced to the olfactory mucosa and olfactory epithelium.
The trace amine-associated receptors (TAAR) for which humans have 7 genes (TAAR1, TAAR2, TAAR3, TAAR5, TAAR6, TAAR8, TAAR9) and 2 pseudogenes (TAAR4P, TAAR7P) are found in the olfactory epithelium whereas androstadienone receptors are found in the olfactory mucosa.  As suggested by the anatomical location, these receptors interact with the human sense of smell and the olfactory system
More information: http://en.wikipedia.org/wiki/Pheromone#Humans

Do you know of any specific studies on the matter?

You could take a look at http://www.ncbi.nlm.nih.gov/pubmed/9515961

While the result of the study you cite is compelling, my understanding is that the pheromone(s) and receptor(s) mediating that response have yet to be identified. As such, the jury is still out as to whether the menstrual cycle effect is really due to a pheromone response. Also, although it is true that we have TAARs, they may not be used as pheromone receptors in humans. I do not think that there is a consensus as to whether human pheromones exist.

I think that this is the most accurate information that we currently have available. As this is a science, it can be expected that results will change as new discoveries are made and old discoveries are disproven.
As far as I know, there currently isn't much evidence disproving the theory and at least some that suggests human pheromone existence, so controversy at the moment is more doubt in limited evidence than two contradicting theories

@arcyqwerty: The major evidence against human pheromone reception is that humans anatomically lack the accessory olfactory system (vomeronasal organ, etc.) that is used by other animals to detect pheromones. Is that strong evidence? That's up for debate, but I don't think either side has much stronger evidence. For what it's worth, I am inclined to believe that there are human pheromones. But that is bald opinion. The question asked for current consensus, and I stand by the statement that there is none.

Wondering if that simply boils down to the sensation of smell. So that pheromones smell good/bad (in the case of bad genetic compatibility).

This article details how man can smell ovulation thanks to testosterone: https://journals.sagepub.com/doi/full/10.1177/0956797609357733 It is also one of the base papers for one of the answers here.

Well... I'm excited that my first contribution to this site will include a study of strippers!
http://whywereason.wordpress.com/2011/06/29/menstruation-attraction-why-females-shouldnt-flirt-while-menstruating-and-why-bears-can-smell-the-menstruation/
I have read other, more detailed analysis of this (and similar) research that suggests strongly that our noses are heavily involved in all sorts of entirely sub-conscious decision making, as it pertains specifically to sexual selection.

A blind test of tshirt scents alledged that people prefer tshirts with complementing immune systems for reproduction.


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

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

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

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

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

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

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

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

Unrooted by default? Which algorithms are you talking about?

Trees can not be distinguished from each other because every unrooted tree can be made rooted by adding a root and vice versa by deletion.
If you mean dendrograms... the root is always the one node where the distance to each leaf is the same. But I don't know of an unrooted dendrogram.

Yes, but how would I be able to tell whether or not a tree has a root
Like how to tell between an unrooted tree and a midpoint rooted tree?

If you really mean trees. They are not distinguishable.

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


text:
What are limiting factors for intron length?
For predicting genes from a sequenced genome, you need to set a maximum intron length. How long can introns get in animals? Is there some limit?

I don't know about the maximum length, but I suppose that for the minimal size, the mechanical properties of the DNA are the limiting factors. Under a certain size, DNA is simply too stiff to form the loops required during splicing.

I think the above values (500-750 kb) are wrong.  http://www.bioinfo.de/isb/2004040032/ shows that most introns are less than about 10 kb (and personal experience in Drosophila confirms that - I've rarely seen an intron bigger than about 5 kb).  There are some very large ones, but since it's nearly impossible to detect the splicing reaction, particularly if they're very large, it's not clear whether they are spliced as one BIG intron or are instead recursively spliced (in little steps, as in http://www.genetics.org/content/170/2/661.full.pdf+html).  I think its safe to say that we don't know how long the biggest introns can be.

If you examine the human genome ~99% of the introns are under 500 kb. I would assume that a limit between 250 kb - 500 kb is reasonable for gene prediction.  You may incorrectly predict the proper structure of a small number of genes that have these very large introns but this should be a small number.  Furthermore, most popular sequence aligners tend to set an intron length limit between 500 kb and 750 kb.
Just keep in mind that you may increase the number of false positive introns you detect if you set this limit to high.  Therefore, it may be worthwhile to try a few settings and evaluate the results.
EDIT:
My guess is that larger introns are constrained in mammals for two reasons

They are more difficult for the spliceosome to excise properly.  The spliceosome may have reduced assembly at the proper 5' / 3' splice sites and the branch point site.  There is also a higher chance that there will be cryptic / decoy splice sites witihin the intron.
They increase the time it takes to transcribe the gene


text:
Which sequence characteristics influence the transcription efficiency of T7 polymerase?
The T7 polymerase doesn't transcribe all sequences equally well, the transciption efficiency can vary widely for different sequences. One well known requirement of the T7 polymerase is that the sequence should begin with 2-3 guanines for efficient transcription.
Is there any literature that quantified the effects of the sequence on the transcription efficiency? I'm mostly interested in the effect of the first few nucleotides, as changing other parts of the sequence is usually not possible.

Would be nice to know why some of my T7-promoter plasmids produce ~100 mg protein/L of culture and others maybe 2-3.

Well, the protein expression also depends on the strength of the ribosome binding site.

I'm doing in vitro transcriptions and I'm not translating the RNA anyway, so for me that part doesn't matter

This is not an answer, but a starting point. I found this article Bai L, Santangelo TJ, Wang MD. Single-molecule analysis of RNA polymerase transcription. Annu Rev Biophys Biomol Struct. 2006;35:343-60. that may contain the answer to your question. Lacking that, you may find good starting points in the references section.

By far the most important part is the very beginning of the transcript, especially the positions +1 and +2. The conservered consensus sequence in class III T7 promoters is GGGAGA, any changes in the first two nucleotides severely reduce the transcription efficiency. Changes at positions +3 to +6 have much smaller effects.
Additionally, changes that put many AU base pairs in that region (e.g. GGUUU) seem to affect the trancription efficiency negatively.
Other sequences that are problematic anywhere, not only in the beginning are long stretches of uridines or adenines. Sequences with eight or more uridines or adenines can cause the polymerase to slip, which results in transcripts with more uridines or adenines than in the template.
These characteristics are detailed in the paper from Milligan and Uhlenbeck from 1989.

Milligan, J. F. & Uhlenbeck, O. C. Synthesis of small RNAs using T7 RNA polymerase. Meth. Enzymol. 180, 51–62 (1989).


text:
Is there any convincing evidence for the existence of nanobacteria?
The existence of nanometer scale microorganisms has been proposed and used explain several phenomena  including morphological structures in a martian meteorite (ALH 84001) and implication in the formation of kidney stones (1).
Is there now any consensus in the biology community whether these are biotic or abiotic in nature?
(1) Kajander EO, Ciftcioglu N (1998) Nanobacteria: An alternative mechanism for pathogenic intra- and extracellular calcification and stone formation. Proc Natl Acad Sci U S A 95: 8274–8279

There are two convincing papers (1,2) arguing that the observed "nanobacteria" are in fact mineral/protein complexes and not any living organisms.
In (1) Martel and Young created very similar looking particles from calcium carbonate in vitro. The calcium carbonate precipitations can even look similar to dividing cells:

After 5 days of serum incubation, a white precipitate formed and
  adhered on the bottom of each flask. When examined by scanning
  electron microscopy (SEM) and transmission electron microscopy (TEM),
  the precipitate was seen to consist of particles with a size ≈500 nm
  (Fig. 1 A, B, and E), an observation consistent with normal bacterial
  morphology, which peaks at one particular size specific to the strain
  on hand. Some forms resembled cells undergoing division (Fig. 1 D,
  arrow).

One strong argument against those nanobacteria being living organisms is that they don't seem to contain any nucleic acids. Of course non-DNA based organisms are theoretically feasible, but that would be an absolutely extraordinary discovery.
The autors of (1) also observed these particles after filtration through a 0.1um filter. They argue that this size could not be sufficient to contain all the cellular machinery bacteria need:

Because an earlier workshop commissioned by the NAS had suggested that
  the minimal cellular size of life on Earth must exceed 200 nm in
  diameter to harbor the cellular machinery based on DNA replication
  (19), it is unlikely that nanobacteria represent living entities
  unless they contain some other type of replicating mechanism.

(1) Martel J and Young JD, Purported nanobacteria in human blood as calcium carbonate nanoparticles. PNAS 2008, 105(14):5549-54
(2) Raoult D. et al., Nanobacteria Are Mineralo Fetuin Complexes, PLoS Pathog. 2008, 4(2): e41.

It would be very illustrative if you could post the actual figures shown in these papers.


text:
How can you identify if a person is homozygous for a certain allele?
I've been thinking about starting a small private research project. In this project I need to find out whether a person is homozygous for a certain allele. The reason for this is that I'm really trying to find out whether has a mutation that makes him/her not produce this chemokine. Thus I need to identify in an easy way which people are "normal", which people have the mutation in one chromosome, and which people have the mutation in both. About 1% of the population have the mutation in both chromosomes.
A primer would AFAIK not be able to differ between the heterozygous and the "normal" people.
Do anyone know how to approach this?

you could use a test cross but that might get complicated :)

I don't think I have time to breed humans ;)

You could use a 3-primers strategy, where you have a common primer, one that only amplifies the mutant allele and one that only amplifies the wt. This may or may not be feasable depending on the type of mutation.
DNA from het people will amplify both product, while homozygous will only amplify one (wt or mutant).
Of course then, linking the mutation of a gene to the lack of a chemokine is not necessarily straightforward.
To be honest, however, I would use a different approach: take people who don't produce the chemokine, and people who do (simply by dosing the chemokine in the blood). Now sequence the gene in the two groups and look for mutations.

Yeah, that's a smart approach. But I don't think (= don't know if) the nonfunctional gene will look the same in all individuals. Since it's a missense/nonsense mutation, it probably doesn't have a clear characteristic. Anyway, good point! +1

@Pedery: well, then wouldn't the sequencing be the best strategy in that case? Do you have any information whatsoever on the mutation or its product?

Not at the moment. Right now I'm trying to get an overview. Sequencing would not really be an option cause I want to make this into a super easy testing-kit, one that would need no expensive machinery and would get a test result from a mere droplet of blood. But indeed, your answer was helpful. If I don't get any better answers, I'll mark yours as accepted.

Btw, what would be the purpose of the common primer? If you have the two other primers, they will still tell you all the information you need, right?

Nope, you need two primers for doing a PCR amplification. If you only have a primer that goes on the mutant allele and one that goes on the wt you will have no way of amplifying anything, as the two will never bind on the same DNA.

Ah, got it. I was thinking in sets of two primers per allele as one primer.


text:
Why is there now only one Salmonella species?
Once upon a time, I chanced upon an old microbiology book that detailed the rather colorful world of enterobacteria. Salmonella in particular stood out, as it seemed there were a lot of species: typhi / typhosa, paratyphi, gallinarum, typhimurium, choleraesuis, and quite a bunch of others that I have now forgotten.
Flipping through a newer book, it now seems that all of these "species" were collated under choleraesuis (and now more recently enterica), with all those species being demoted to "strains" (or maybe I should use the current term of art, "serovar"). Unfortunately, the book didn't give much in the way of explaining about this merger.
So, why is there now only S. enterica? If "S. typhi" is a mere serovar, how come the species name is still used in the literature?

Simply put, old habits die hard; physicians and other medical personnel have grown up with the old species designations so will continue to use them.  This is somewhat the reverse of the case with E. coli, where 80-90% of the genome is variable across strains.
Lin-Hui gives a brief history, where strains identified early were given specific names within Salmonella like typhi, typhimurium, panama, etc.  In 1966 it was proposed to lump serovars together among three species S.
choleraesuis, S. typhosa, and S. kauffmannii.  Shortly thereafter in 1972 with the advent of DNA techniques, it was shown they really should be all piled together.  There is one exception, Salmonella choleraesuis subsp. bongori, separated in 1989.

Nice survey article! (Even the switch from choleraesuis to enterica is discussed.) Thanks for this. I believe you intended "typhosa" instead of "thphosa", though.

@J.M. probably...it's misspelled in the review though :/

Just open Wikipedia, your question is answered there:

Initially, each Salmonella species was named according to clinical
  considerations,[15] e.g., Salmonella typhi-murium (mouse typhoid
  fever), S. cholerae-suis (hog cholera). After it was recognized that
  host specificity did not exist for many species, new strains (or
  serovar, short for serological variants) received species names
  according to the location at which the new strain was isolated. Later,
  molecular findings led to the hypothesis that Salmonella consisted of
  only one species,[16] S. enterica, and the serovar were classified
  into six groups,[17] two of which are medically relevant. But as this
  now formalized nomenclature[18][19] is not in harmony with the
  traditional usage familiar to specialists in microbiology and
  infectologists, the traditional nomenclature is common. Currently,
  there are two recognized species: S. enterica and S. bongori, with six
  main subspecies: enterica (I), salamae (II), arizonae (IIIa),
  diarizonae (IIIb), houtenae (IV), and indica (VI).[20] Historically,
  serotype (V) was bongori, which is now considered its own species. The
  serovar classification of Salmonella is based on the Kauffman-White
  classification scheme that permits serological varieties to be
  differentiated from each other. Newer methods for Salmonella typing
  and subtyping include genome-based methods such as pulsed field gel
  electrophoresis (PFGE), Multiple Loci VNTR Analysis (MLVA), Multilocus
  sequence typing (MLST) and (multiplex-) PCR-based methods.[21]

Beautiful. Now, what were these "molecular findings" that were instrumental in collating all those Salmonella "species" (I don't have access to the journal article linked to)? And, why is "S. typhi" still used instead of "S. enterica serovar typhi"?