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Can protein structure be determined by X-Ray Diffraction in a single image?
I'm reading about the use of x-ray crystallography to determine protein structure. According to my book, data is collected at 30-360 angles (dependent on the symmetry of the protein). An illustration is given with concentric rings labelled with distances - the further out the points are, the higher the resolution.
Is the image a composite (where the angle of the point point from the centre is equivalent to the angle of the reading) or is a separate image taken at each angle? Are there any other reasons why more images would be required?
Thanks.

I can't answer on the details, but as far as I understand it they take multiple images while changing the angle.

That's right. I was wondering if the images could be superimposed somehow, since that was not very clear to me.

It is not clear from what you write whether you are familiar with the process of analysis of X-ray diffraction. Is it clear to you that the diffraction pattern has to be mathematically analysed to get the final "3D picture" of the crystal?

I have to agree with the answers below, it's best to collect light diffraction from the crystal at multiple exposures within the 0 - 180 degree range.

You cannot solve a structure with a single frame, even with perfect diffraction.
The reason you need images over a large swath of angles is because the diffraction pattern is also in three dimensions, in the so-called "reciprocal space".  At minimum, a 180° rotation of the crystal is needed to sweep the entire reciprocal space sphere with the plane of detection, though symmetry in the crystal structure can reduce this further (some of my proteins only required 90°, I think hexagonal unit cells with 6-fold symmetry can live with 30°).  Because crystals are real things (read: non-ideal, doubly so for protein crystals), the sweep is extended to gain some redundancy.

Not by analysing a single protein.  There is work with x-ray lasers.
You have to take a simultaneous image of millions of proteins and use that to get a structure.  It's not quite prime time.  People are also doing this with electron beams in electron microscopes.
These methods will reconstruct 3D models of the molecules, sometimes in states which cannot be obtained from crystallography.  Examples being the structure of the many megadalton nuclear pore complex, and the f-actin fiber.   The classic study is  3d model of bacteriorhodopsin, the first membrane protein structure which was at molecular resolution (this was a crystalline sample though).
While in principle, it sounds much simpler - get a pure sample of your protein, or complex and freeze it down and zap it with an Xray or Electron beam, its a lot more work to reconstruct the image and can take as long or longer than getting an x-ray structure.  The resolution is also usually poor as the crystal will reinforce coherence, that is all the proteins are aligned in the same way and have close to the same 3d shape in a crystal.

Thanks for your answer. I didn't mention it in my question, but my book does say that a vast number of crystalised proteins are required, and that simultaneous diffraction amplifies the signal. What I was trying to ask is if a type of crystalised protein conformation can be determined from a single image or if many different images would need to be produced for each angle of rotation. From your answer it sounds as if all the information required is stored on one image - is this correct?

I'm confused by your answer, what do you mean by "it's not quite prime time"? Electron microscopic images also don't provide nearly the resolution you can achieve with X-ray crystallography.

@MadScientist I think the implication is that superimposed images are also used in EM to reconstruct the structure. The case is somewhat different of course since EM in general doesn’t use crystallised structures so that the proteins in the image don’t all have the same orientation, which is crucial.

CryoEM also doesn't use any X-ray beams either.

I think that while they are working very hard, the X-ray laser and cryo EM reconstruction doesn't yield as consistant a set of results as crystallography.  when the proteins just lay on a surface, they can be shape distorted, and so recombining the images makes can be difficult to interpret.

@Mad Scientist comment started running long, edited answer..

Likely not. While one can get excellent diffraction data from a high quality crystal, it would be extremely difficult to solve the phase problem. The extra angles will help constraint the solutions.

Can protein structure be determined by X-Ray Diffraction in a single image?

Yes. Using a technique called Laue diffraction, it is possible to obtain sufficient data from a single image to solve a protein crystal structure. One example is the time-resolved study of carbonmonoxymyoglobin dissociating by photolysis (DOI: 10.1107/S090904959501661X). This is not the standard single-wavelength technique that is usually used, but uses "white" X-rays with a range of wavelengths available only at synchrotron beams. For example, the BioCARS user facility provides infrastructure for time-resolved crystallography. It is also used in "diffraction before destruction" when working with free electron lasers, see for example Nature Methods 8, page 283 (2011).
The remainder of the answer is about conventional single-wavelength crystallography.

Is the image a composite (where the angle of the point point from the centre is equivalent to the angle of the reading) or is a separate image taken at each angle

The desired structural information (3D electron density in real space) is a Fourier transform of the diffraction data (3D reciprocal space). The word "image" is jargon for a single diffraction image, i.e. the diffraction spots you observe when you point an X-ray beam at a crystal in a certain orientation. Using a different orientation, you get more data (and also measure some spots multiple times).

Are there any other reasons why more images would be required?

The more diffraction images are collected, the higher the completeness and redundancy of the data. Completeness refers to having measured every diffraction spot at least once. Redundancy refers to how often on average a spot was measured, and increasing redundancy increases the quality of the measurement through averaging.

I accept this answers the specific question "Is it possible?", but this was rather a proof of concept. Do you know whether this has been adopted to any extent — found use in circumstances where it was not possible to obtain more images? Also, could you cite the original reference. Researchgate is not a journal.

This is used routinely in time-resolved crystallography and in "diffraction before destruction" when working with free electron lasers XFEL, Nature Methods 8, page 283 (2011)

Thanks. You should add this to your answer and I will upvote it. Unfortunately this list is dominated by what I call “furry animal” biologists. Nothing wrong with that, I suppose, but the effort put into factual hard scientific answers is seldom rewarded in Brownie points.

@David Yes, I was visiting from StackExchange Chemistry. There are question in Biochemistry that could go either way, just as there are questions that fit in physics or chemistry, but will get a different type of answer (or none). Not that there is anything wrong with organismal biology.

As long as a sufficient amount of the reciprocal space has been collected, then it is of course possible. How the phase problem is handled when doing Laue diffraction I simply don't know, but I would assume Molecular replacement or Experimental phases can be used?

@JeppeNielsen Often, the structure is solved by Fourier difference, i.e. most of the unit cell is identical to a known structure, except for changes in the ligand binding site. With free electron lasers, one could in principle build a phase-sensitive detector and measure the phases directly.

@KarstenTheis, what you call fourier difference is equivalent to Molecular replacement phasing. I don't know how one would create a phase sensitive detector for an X-ray Free Electron Laser (XFEL) when one cannot do so for synchrotron based X-rays. The major difference between the two light sources is the peak photon flux and pulse duration. Are you aware that the electrons are separated from the x-rays prior to the diffraction experiment, thus you have to determine both phase and intensity of the diffracted x-rays, not the charged electrons.

@JeppeNielsen Molecular replacement requires a rotation and translation search, often with Patterson methods. Once the orientation of molecules in the unknown structure is known, Fourier difference methods are used to adjust the model.

@JeppeNielsen The difference between XFEL and synchrotron radiation is, as far as I understand, that XFEL is coherent (all the same phase) and synchrotron radiation is not (multiple electrons adding up to create the X-rays, not coherent).


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What determines a successful protein expression in E. coli?
Some proteins express well in a heterologous host; others- don't. A few requirements are known to determine the protein expression, like a strong promoter (like T7) for transcription and a strong ribosome binding site for translation. I am working with a protein, which consists of 2 subunits - alpha and beta. Both of them are on a plasmid with T7 promoter in front of the beta subunit (i.e. the construct is T7 promoter, CDS for beta subunit, CDS for the alpha subunit). The beta subunit expresses well, but the alpha doesn't. Do you thing this has something to do with the local environment (promoters, RBSs, etc) and how much does it depend on that? How can I increase the protein expression?

Some aspects are not entirely clear to me. The two subunits are expresses on seperate plasmids, or? And everything in the plasmids is the same except for the actual protein sequence?

The two subunits are on the same plasmid. The plasmid sequence starts with T7 promoter, RBS1, subunit beta, RBS2, subunit alpha. The RBS sequences are a little bit different, so I was thinking of making them the same, but this shouldn't have a great effect on the protein expression. (they are very close to the consensus sequence).

I found a very nice paper: Designing Genes for Successful Protein Expression, which covers most factors that determine protein expression. I post parts of it, because I am sure it will be useful to some of you.

Translation can be controlled at the level of initiation and elongation. Initiation of translation is primarily dependent on the sequence of the ribosome binding site (RBS) and early mRNA secondary structure. Other determinants of protein expression are less well understood but equally potent.
1. Initiation of translation
A key component affecting initiation of translation in prokaryotes is the RBS that occurs between 5 and 15 bases upstream of the open reading frame (ORF) AUG start codon. Binding of the ribosome to the Shine–Dalgarno (SD) sequence within the RBS localizes the ribosome to the initiation codon... Affinity of the RBS for the ribosome is a critical factor controlling the efficiency with which new polypeptide chains are initiated. This interaction is in competition with possible base-pairing interactions involving the RBS region that may form within the mRNA itself. Thus, SD sequences with weaker base pairing to the ribosome are more susceptible to interference from mRNA structure. However, some experiments suggest that SD sequences with too strong affinity can be deleterious, particularly at lowering temperatures, by stalling initial elongation. Also critical is the distance between the RBS and the start codon with 5-7 bases from the consensus SD AGGAGG being optimal.
Numerous lines of evidence suggest that the initial 15–25 codons of the ORF deserve special consideration in gene optimization. Studies have shown that the impact of rare codons on translation rate is particularly strong in these first codons, for expression in both Escherichia coli and Saccharomyces cerevisiae. In E. coli, peptidyl-tRNA drop-off during translation of the initial codons appears to be accentuated by the presence of rare NGG codons. These effects appear to be independent of local mRNA secondary structure. It is also true that expression may be recovered by 5' sequence replacement even for sequences that do not show especially strong mRNA structure or contain rare codons or other obvious deleterious elements in this region.
2. Codon bias
The second way in which host codon frequencies can be used is to match the host codon frequencies in the designed gene. This can be done simply by choosing each codon with a probability that matches the host codon frequency...Using sets of genes broadly varied in gene design features, Welch et al. found that variation in synonymous codon usage frequencies had a profound effect on the amount of protein produced in E. coli, independent of local 5’ sequence effects. Variation of at least two orders of magnitude in expression was seen due to substitution beyond the initial 15 codons of the ORF. This variation was strongly correlated with the global codon usage frequencies of the genes, although the codon frequencies found in the highest expressed variants did not correspond to those found in the genome or in highly expressed endogenous genes of E. coli. Multivariate analysis showed that the frequencies of specific codons for about six amino acids could predict the observed differences in expression. It is not clear what the biochemical basis is for this correlation.
3. mRNA structure and translational elongation
While much evidence suggests that mRNA structure can interfere with translational initiation in both prokaryotes and eukaryotes, the effects of structure on elongation are less well understood. This in part may be due to intrinsic helicase activity of ribosomes, which allows translation through even very strong hairpins and may preclude many structures from limiting the translation rate in either prokaryotes or eukaryotes. Perhaps more importantly, mRNA structure is difficult to predict, particularly for actively translated messages which are in continuous flux between various folded and unfolded states.
4. Protein-specific factors providing additional complexity
The protein may be particularly unstable in the host, especially if it is poorly folded due to inherent instability, lack of sufficient prosthetic factors, or improper post-translational modification... Expression of secreted and membrane proteins may be limited by mechanisms for directing these proteins to the membrane. It is even possible that the protein amino acid sequence may limit translational efficiency. For example, proline is thought to be slowly translated in E. coli, regardless of which codon is used.
Expression of the protein may be toxic to the cell leading to instability of the expression vector or host suppression of protein synthesis...A common strategy to reduce toxicity is to lower expression to tolerable levels. Promoters varied in strength can be valuable tools for finding an optimal expression rate for maximal yield...One potential way to avoid toxicity of some proteins is to direct expression to the periplasm or media. This may be accomplished by N-terminal fusion of a secretion signal sequence.

For more information, please read the whole paper. I also recommend reading Design parameters to control synthetic gene expression in Escherichia coli.

Thank you for the links! I've used DNA 2.0 for codon optimization & gene synthesis before and have had good results - I'm definitely interested in reading their publications.

@Amy: Yeah, part of the paper is about the features of DNA2.0, which I find very useful. I am a new user and still getting used to their software, but it looks pretty nice so far.

One important aspect when expressing a protein from a different organism in E. coli is that the codon usage of the original organism is likely different from the codon usage of the E. coli used for expression (Codon usage bias).
Though the genetic code is degenerate, not all codons are equal. They might encode for the same amino acid, but organisms tend to favor specific codons over others and the tRNAs for those codons tend to be present in different concentrations. When you now have a lot of codons in your sequence that are very rare in E. coli, the expression will suffer because the tRNA for those codons is not present in high enough concentrations to sustain the translation.
There are two ways of compensating for that, either you get your E. coli to produce more of the rare tRNAs, or you optimize your sequence to use different codons. For the first solution you can buy certain E. coli strains that contain plasmids encoding the tRNAs that are rare in E. coli, one of those strains is e.g. the Rosetta BL21 strain.
For optimizing the codon usage there are several companies offering to synthesize optimized sequences. There are also tools available online, but I have no direct experience with those.
The optimization is also not as easy as just using the most prevalent codon every time, it has been shown that optimizing the codons to match the translation speed in the original organism can enhance the expression yields. For some proteins, pause sites during the translation might be necessary to ensure proper folding. If the protein doesn't fold correctly it will be quickly degraded and your yield will suffer. This is described in the article "Heterologous Protein Expression Is Enhanced by Harmonizing the Codon Usage Frequencies of the Target Gene with those of the Expression Host" from Angov et al.
You'll find a nice overview about the whole topic in the article "You're one in a googol: optimizing genes for protein expression" from Welsh et al.

Thanks, Mad Scientist! Yes, I have thought about codon optimization and this is definitely on my to do list. Sorry I forgot to mention it. DNA 2.0 has a software which will convert your sequence into optimized one. I have also heard about Rosetta BL21, but I have never used it. We might also want to stick to our own strain, because it is the heterologous producer of of or drug of interest. I am a little concerned about codon optimization, precisely because of misfolding but I thing we should do it.

But this still doesn't explain why the other subunit expresses better, as well as other proteins from the same native organism are well expressed in E. coli. That's why I think it is more of a question about the control elements of transcription and translation, rather than rare codon usage.

@GerganaVandova You should compare the codon usage of your two subunit, it might be just by chance that one uses more rare codons than the other one.

I'm attending a talk by Claes Gustafsson from DNA2.0 - and i guess we knew this already - that codon usage is helpful, but doesn't always work.  they have a proprietary method which is more reliable...

Have you tried putting your two genes on two separate plasmids (with different origins of replication and antibiotic selection, of course) and co-expressing that way? If the first of the two subunits is expressing well and the second isn't, that's probably because the ribosomes are falling off the mRNA before fully transcribing the second gene. Are the genes eukaryotic in origin? I think it is difficult to "trick" E. coli ribosomes into treating heterologous sequences as an operon, but maybe someone with more knowledge of prokaryotic translation could help.
Actually, just adding a second T7 promoter in front of gene 2 would probably help quite a bit:

Genes are transcribed either from individual promoters or from a single promoter, leading to a long polycistronic mRNA. Polycistronic expression has been reported to lead to lower expression of the more downstream encoded protein, which can be exploited to influence the stoichiometry of a protein complex (9). Several-fold higher expression with individual promoters compared to polycistronic transcription has been reported (10). (Source)

I do know from personal experience that co-expression using two plasmids can work very well with the right genes!

Thanks for your answer. I like the idea of adding additional T7 promoter, although having repetitive sequences might lead to other future problems (I don't want to go into details). But I work with genes which are 3 times larger in size and one promoter for the 2 subunits works well. Perhaps it is very sequence specific.

Mad Scientist covered potential codon-bias issues (which can be ameliorated with Rosettas), but more generally, I've seen tremendous variability in expression rates between different vectors (pET-28 vs pET-24) without any apparent reason.  Our lab has had tremendous success with IPTG-inducible vectors (going from pBC-SK to pET-24 increased expression 50-fold).
Beyond expression, the protein can be lost when preparing the cell-free extract.  It may not be in the cell pellet if exported courtesy of a signal peptide, or may be discarded in the "debris" pellet when spinning down lysed cells if it's poorly soluble.  I've heard a theory that solubility issues may be exaggerated by vectors that saturate the export machinery, which can both impede synthesis and/or be so effective they just cause precipitation.  The pET System Manual suggests longer (overnight) expression times at lower (15-20 °C) can help solubility issues.  Whatever the reason, engineering away the signal peptide drastically increased expression for many of our proteins.

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What causes the development of antibiotic-resistant strains of bacteria?
I understand bacteria have become resistant to antibiotics due to selection pressures, but how do resistant bacteria process antibiotics when exposed to it, compared to non-resistant bacteria. Also, what research is being conducted to combat bacteria resistant to antibiotics?

I suggested an edit for the title, the original one sounds like all bacteria are antibiotic-resistant.

@J.M. + cancer as well

Bacteria usually gain resistance mechanisms through horizontal gene transfer (such as conjugation and phage infection). The four main mechanisms in which bacteria elude antibiotics are:

Drug Inactivation: For example, E. coli can produce beta-lactamase that inactivate many lactam based antibiotics such as penicillin.
Alteration of Target Site: Mutations in genes encoding for target sites can reduce drug binding affinity. Example: Random mutations in DNA gyrase and topoisomerase IV reduces binding affinity to fluoroquinolone.
Alteration of Metabolic Pathways: Many drugs target certain parts of metabolic pathways by inactivating enzymes or sequestering substrates. Bacteria can use alternative metabolic pathways or find ways to uptake the necessary nutrients from the environment.
Reduced drug accumulation: Caused by reduced drug permeability or the ability of the bacteria to transport drugs out of the cell (for example, tetA encodes for a tetracycline efflux transporter). They can gain these abilities through conjugation, phage infection (transduction), or uptake of environmental DNA (transformation).

Wikipedia has a pretty extensive and comprehensive article on antibiotic resistance. There's also a good list of references. http://en.wikipedia.org/wiki/Antibiotic_resistance

Also, what research is being conducted to combat bacteria resistant to antibiotics?

You've gotten some decent answers for how antibiotic resistance arises, so I thought I'd touch on this a bit. There's three major thrusts to anti-resistance research:

Finding new targets and mechanisms. Essentially, creating new antibiotics that are subtly different enough that they slip past existing resistance mechanisms (usually by tacking a more elaborate side-group onto an existing antibiotic) or an entirely new class that targets a different mechanism to disrupt bacteria. This is essentially a drug development task, and is hard.
Antimicrobial stewardship. This is largely clinical research examining how we use are existing antibiotics. Can we alter the duration/deployment/etc. of a course of antibiotics to minimize the chance of resistance. Can we minimize the number of times we use antibiotics inappropriately, giving rise to resistance with no gain to the patient?
Alternate antimicrobial techniques. Can we use non-antibiotic techniques to prevent bacterial infections in the first place, or treat them when they occur? Antimicrobial surfaces, like copper or silver impregnated surfaces, improved surface cleaning, etc.

The lead question you have answered yourself: bacteria become resistant because of the selection pressure caused by the antibiotic's effective suppression of the original non-resistant bacteria.  Those variants which resist the suppression are selected for as a natural consequence.
How do resistant bacteria process antibiotics?  It depends on the details of the particular antibiotic, and perhaps the kind of resistance.
Take the case of penicillin and related antibiotics, like amoxicillin.  These antibiotics act by inhibiting the formation of a layer of the cell wall which is essential for many kinds of bacteria.  This prevents multiplication of the bacteria and contributes to their distruction. Wikipedia gives some details on the action of these β-lactam antibiotics.
Bacterial resistance to drugs like penicillin usually takes the form of the bacteria producing an enzyme (called β-lactamase) which breaks apart a ring in the drug molecule, disabling it and thus removing its effect on cell wall synthesis.
This resistance has become common because of the widespread use of  penicillin-like drugs and because of transfer of the gene for it between bacteria species as by plasmids.
Researchers managed in the 70s to discover and develop an auxilliary weapon in this war.  This was clavulanic acid, which has a structure partly similar to the penicillins and like them is attacked by the bacterial β-lactamase enzyme.  Unlike the penicilins though, it forms a permanent bond with the enzyme molecule, disabling its activity.  This has led to currently effective drugs like Augmentin which include in the same pill both amoxicillin and its protector, clavulanic acid.

@J.M. and the route taken by bacteria is to evolve/select for beta-lactamases that have activity against carbapenems, like some of the lesser-prevalent (at present) class-D enzymes, thus ensuring a continuing source of funding for our lab.

@J.M. you're eluding to how calling it a "war" is a bit of a lie; reminds me of a guy on TotN talking about the "war on cancer"

Most resistance is acquired by horizontal transfer by various means
Conjugation is the sexual transfer
Transduction is transfer by viruses that integrate into the  genome then when they are activated, they carry bits of the genome attached to their own to a new host
Transformation is the uptake of naked plasmid DNA into a new host
Transfection is similar to transduction and is viral mediated
And of course you have acquired which results from point mutations that alter the cells ability to be harmed by an agent, simple changes like change of the cellular membrane or receptor sites

thanks for your answer and efforts in the community... we always appreciate references to back up the information stated in answers as well as questions... do you thin you could add some links to help others follow your answer (I know some of this may be common knowledge to you, but citations and other sources - even webpages - are better than nothing). Also, you may want to check out the help site on "how to write a good answer" http://biology.stackexchange.com/help/how-to-answer

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Solution based measurement of Solvent-Accessible Surface Area of macromolecules
The Solvent-Accessible Surface Area (SASA) is a valuable metric for looking at protein folding and protein-protein interactions. However, this measurement is typically done by calculating the SASA from a solved (and generally static) structure.
Chemical probes like diazirine and hydroxyl radicals show some bias regarding where they tend to bind. I'm realizing while I'm writing this question that NMR is a perfectly valid method to determine a solution based structure and then calculate the SASA. Similar strategies has also been used to examine structured RNAs. I'm curious about the variety of these methods and how accurate they are.

Determining an NMR structure is a lot of work, it's not something you would do just to get that information. There are ways to identify which residues are solvent accessible by NMR (e.g. SEA-TROSY), but they don't give you the area.

I only know of one method, but here it is.  You create a sphere the diameter of the VdW radius of water, and then 'roll' it along the surface.  I know this as a Richards-Lee surface, wikipedia has another name for it.

This looks complicated, but its not.  you move the probe sphere along the surface of the molecule in the XY plane until it just touches the vdW radius of the protein, keeping the center of the sphere as the surface, all the way around the molecule.  If you like, you can color the surface by the charge of the position too, which is useful for discussing solvent interactions.
Then you translate along the z axis and do another contour until you run out of protein.  Apparently jmol and other packages will do this for you.
Wikipedia references a more mathematical method LCPO, which I am not so familiar with.
Is this accurate?  As usual with such calculations its more of a guess than an answer.  You can do the calculation on any structure or any ensemble of structures (like NMR gives).  It doesn't understand how the molecule might be flexible or dynamic.  If you read up on your physical chemistry you see that proteins breathe and can allow diffusion into the core rather readily.  If I recall right, you can get rather large molecules quenching heme flouresence in hemoglobin at room temperature.
If you are looking to dock 2 proteins, SAS might be more useful.  Its an important piece of information, but not an ultimate answer.  I'm afraid with proteins that doesn't happen so easily.
@bobthejoe asked about SAS for which no structure exists.
This is an extremely difficult thing to even guess at.  The non helpful answer is that the surface of the protein goes as the cube root of the molecular weight of the protein.
By getting a solution of the protein and shooting it in a syhcrotron, you can get a mean radius of gyration pretty easily which will give you an ellipsoidal volume (and surface area) for a protein.  Again most of the particulars would be lost and this could easily be off by 25% for an irregularly shaped protein.  For a regular globular protein it might give an answer similar to the power law above.
I have seen physical chemistry experiments that look for changes in osmotic pressure when the salt concentration in a solution of the protein changes substantially (Adrian Parsegian's work at NIH in the late 80s).
I doubt you will find any of these answers useful as their mean error is going to be very large (20-200%) and also assumes the protein is soluable and amenable to the experimental conditions.
Solvent probes can help too.  For instance exposing the protein to D20 then doing mass spectroscopy on the protein. This is still only going to give you a general idea of how much of the peptide is surface exposed.  Protein structure is still pretty necessary to getting any accurate measurement of SAS I think.

This is the mathematical calculation of SAS. Unfortunately, it doesn't get to the answer of how does one measure SAS sans X-ray/NMR structure.

sorry i missed this.  i have added something about this.  this is difficult to do as you might have guessed.

I don't think SAS is a well-defined observable. (I like the work by Bertil Halle on this and related problems.) You can measure correlates to SAS that have been calculated according to known structures, using something like the rolling sphere algorithm that @shigeta describes.
I think the best-parametrized approach would be differential scanning calorimetry. Because a lot of proteins have had their folding studied with this technique, a DSC curve can be directly converted into changes in polar and nonpolar SAS. The total SAS would be a sum of those two quantities.
The only ready reference I could find for this is here: https://web.archive.org/web/20100228041032/https://www.bio.cmu.edu/courses/03438/LecS00/DSC.html
I think most of the analysis was worked out by Ernesto Freire. Basically, the idea is that surface accessible hydrophobic areas organize more solvent (and have a larger entropic footprint) than the polar areas. Since a DSC trace (for a two-state folder) can be deconvoluted into $\Delta$H and $\Delta$S components, it can be mapped onto $\Delta$ASA$_{folding}$. I wouldn't trust this for anything that was non-two-state or that bound cofactors.

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Troubleshooting bioconjugates migration in a SDS-PAGE gel?
We do a lot of bioconjugation chemistry (click chemistry in particular but also NHS and Maleimide chemistries). Our method to valid the conjugation reactions have been to use SDS-PAGE gels followed with densitometry. However one particular challenge with such analyses is that proteins conjugated with non-proteins like DNA or PEG migrate with an inconsistent MW and also in a smear making it hard to isolate the relevant bands. I assume that this has to do with having a poly-disperse radius of gyration. Are there tricks that I can do to get tighter bands?

where would the poly-disperse radius of gyration arise from if the molecules are coated with SDS?

@Ben, my hypothesis: SDS disrupts hydrophobic interactions as a surfactant. However, something like DNA has purely enthalpic interactions and will not undergo denaturation or be evenly coated by SDS. This is the underlying roots of my problems.

The first step in troubleshooting is to run controls - run lanes with your protein input alone, and the conjugate alone (you may need to play with the type/percentage of the gel if the conjugate molecule is much smaller than your target protein - attaching biotin to a 50 kDa protein, for example) to see if the smear shows up. Depending on the type of conjugation you're using, the number of acceptor sites on the substrate, and the time and conditions of the conjugation reaction, you may be adding a highly variable number of conjugate molecules to your substrate. For example, when I first started working with adding fluorophores like Alexa and DyLight dyes to antibodies, the fluor/protein ratio was highly variable from reaction to reaction and between antibodies in the same reaction.
This is most likely what is happening in your case - the number of conjugate molecules being added to each substrate molecule is highly variable. If you haven't already, I'd suggest getting in touch with your conjugation kit vendor's tech support, as they likely encounter this problem all the time. You may need to change the type of conjugation chemistry, get a higher-quality conjugate, or purify the substrate before and/or after the reaction. One of the solutions to my antibody labeling problem I mentioned above was to purify the reaction products through a size-exclusion column. This had the dual effect of separating out the un-reacted antibody and fluorophore, and allowing us to pick different fractions of the conjugated product and decide which were best for our needs.

+1 Thanks! Issues with the suggestions so far. a) we're not conjugating a protein to another protein so standard commassie staining will not stain the conjugate molecule. b) we are the vendor since everything is done inhouse. c) While we can easily separate the reacted molecules from the unreacted, this doesn't answer the problem of quantifying the extent of the reaction.


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What is the highest competency possible for E coli?
I am looking to find a highly competent E coli strain.  I am making a library of a ~6.6kb plasmid and I am not getting high enough efficiency.  Does anyone have a suggestion of a strain/protocol with ultrahigh efficiency (>1010).

I have heard that Epi300 electrocompetent cells (1) from Epicentre are very efficient: > 1 x 1010 cfu/µg of pUC19. Dan Gibson used them in his paper for the synthesis of the mitochondrial genome (2). We were also thinking of using them for our assemblies, but they are pretty expensive.
1. TransforMax™ EPI300™ Electrocompetent E. coli
2. Chemical synthesis of the mouse mitochondrial genome, Gibson et. al.

still this is the better answer - sometimes you have to pay a bit more if you are on the cutting edge.  chemically competent cells are usually not as efficient.


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Which part of oranges contain fiber?
I think it's pith, but here (non-english country) many chef suggest orange's vascular veins which look "fibrous" and have similar translated word with "fiber" itself.
Please enlighten me which?

Cellulose is one of the most common sources of fiber in the nutritional sense. Because oranges are plants, their cells have cell walls, made out of cellulose, so some of the fiber in an orange is surrounding each individual cell. Both vascular cells and pith cells tend to have particularly thick cell walls, so they are probably higher in fiber (this article also suggests that the pith is particularly high in fiber - vascular tissue is probably not mentioned because it is less prominent).


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What to look for when buying a light microscope?
I am a science lover and want to buy a microscope to explore things around me. Like studying cell structures, microorganisms, blood and plants. It would be great if there is way to photograph slides. Just curious to know and explore things around me.
I want to know which features I should consider while buying it?

...and your question is?

To clarify how you could edit your question to make it more suitable for the site, we'd be happier to answer a question asking for an explanation of which features to look out for when purchasing a microscope.

Light. Literally. When I got a microscope as a kid, it was supposed to be lighted with a mirror, which made it hard to find a good setting sometimes. It  was the most frustrating thing about it (for me as a kid).

honestly, save your $,  build your own using parts from Edmonds. Unless your super $$$ not gonna get a very good. this isn't like toyota vs hundae. There are only two manufacturers of microscopes, the prius type (ebay) and the lamborgeini type (zeiss, olympus and leicA). try to get a higher end objective and build your own scope. Optics and light paths are good skills to have too. but i default to @guest below, nice presentation.

Well it depends on the application. First you have to determine what it is that you want to view under the microscope e.g. an organism such as Drosophila or you want to see microscope slides? In the first case you need to by a dissecting microscope, otherwise you need to by an upright light microscope with the appropriate objective lenses, again depending on the application. If you are viewing cells in a flask/dish/plate in a cell culture setting or if viewing a haemocytometer (although not necessarily in the case of haemocytometer), you need to buy an inverted light microscope with phase contrast since you wouldn't be able to view the cells very well with an upright microscope since the goal is to reduce the distance between the objective and the sample for a better view of the samples in cases of dishes or plates. Again the choice of equipment depends on the application. But as correctly said above you need to clarify what you mean in the question.

My advice is always the same: "first microscope? keep it simple, keep it cheap, but nevertheless: buy something usable".
This Olympus GB is, IMHO, "something simple/cheap/usable":

These can be bought here second-hand for anything between € 50 and € 100. It's an uncomplicated stand, easy to overhaull yourself. It permits the use of any optics you would like, it has a nice Abbé-condenser N.A. 1.25. It's heavy,  sturdy and reliable.
These are some of the microscopes I bought second-hand over the last few years. None of those was more than 80 Euro's/US$, except the Diapan (lower right). All were in reasonable condition and ready to use, except one. Notice the build-IN 6V-15W illuminator in the Zeiss GF stand (lower middle), versus the build-ON 12V-100W HAL illuminator in the Reichert Diapan (lower right). See the text:

And if you're handy and not affraid to work it can be even cheaper. This one was for free and it turned out to be one of the best stands I ever had:

Perhaps this can help: it's a (partial) checklist on what to look for when buying a second-hand microscope.
I don't have absolute "no-no's": many problems can be solved, but sometimes it takes a lot of effort, time and cursing...
Sometimes a stand is junk, but it has excellent optics. Or the other way around.
You should take into account that having a microscope (seriously!) serviced/repaired is usually very expensive: a company here, known for their high quality work and their very reasonable prices, asks for a complete overhaul of a microscope genre Zeiss Standard 14/16 a fixed price of slightly more than 700 Euro, spare parts not included... On request they do it on an hourly basis as well: 120 Euro/hour. They do an excellent job, but well... it's not cheap.
1. A first glimpse
Brief examination of a second-hand microscope would take half an hour, up to an hour, depending on how sophisticated it is and on how experienced you are.
You're allowed to be exited and thrilled, but don't loose your critical sense! It's only a microscope. Plenty of those and there are far more important things in life.

Microscope complete, all parts present, all same brand and same
generation (do your homework!). All knobs, screws, gears... present as far as one can see
Suitcase present and in reasonable condition, key present if one is needed, factory certificate present, user manual present
Microscope reasonably clean, no excessive dirt, no severe traces of
corrosion or fungus (if fungus is present, even in very small
amounts: carefully check all optics!). No excessive traces of extremely heavy use. Check aperture diaphragma dial, field diaphragma dial, surface of the stage, x- and y- dials of the stage if present, the surroundings of the focusing knobs, the nosepiece...

Spider webs and dirt on a vintage microscope, stored for decades on an attic. Turned out fine:

Microscope doesn't spread strange smells: fungus, dish soap, cleaning naphta, white spirit, xylene, ethyl alcohol, hexane, greases and/or oils, "creeping oil" such as WD40 or CRC... A microscopes "treated" with creeping oil is about the only absolute no-no I have: these oils do what they're supposed to do: creeping in everywhere, including in the optics
Microscope doesn't show any trace of a fall or a punch. If that kind of damage is present, decent repair should be demonstrated beyond any doubt by official documents only, excluding hearsay: repair costs estimates, invoices, correspondence between owner and insurance company if applicable etc... (observe serial numbers in those documents. They're there for a reason!)

Damage due to a drop in a Zeiss Standard microscope. A cleaning lady dropped it accidently, while cleaning a college lab. Accident and repair were well documented. Replacing it would have been less expensive, but the insurance company insisted on a repair, which was done in the Zeiss factory. Nearly all parts were replaced, except for the base of the stand, as that was considered "cosmetical damage" only. The microscope was equipped with high end optics and the price was modest. A steal:

All adjustments (coarse and fine focussing, nosepiece, condenser height adjustment, aperture diaphragma, field diaphragma, movements of the stage, eventualy present internal magnification changers, Bertrand lenses, polarizers, ... work well over their entire range. Use your senses: sight, hearing and, most important: your tactile sense. Be very aware of "strange things" when using the settings: scratching noises/feel, ticking noises/feel and so on. Might be a minor problem, such as a small lump of dried out grease. Or might be a broken/stripped tooth gear... Very stiff controls, even up to the point of being completely "frozen up" are usually not that big a problem. However, keep in mind it might be necessary to dismantle the microscope completely to solve the problem. That means: dismanteling, cleaning and degraising, regreasing using the right lubricants in the right amounts at the right places, reassembeling and alligning the stand. It's not rocket science, but it is a lot of work! Pay attention to the leafs of iris diaphragms: these should always be dry, and free of grease or oil

Dismanteled coarse and fine focusing controls of a Zeiss Standard. Notice the ingenious construction using a planetary gear:

2. A closer look
3. Microscope under the microscope
Second-hand microscope buyer's best friend: a phase telescope:

"Minor" optics problems...
Seeped in immersion oil and degraded lens kit in a Hensoldt 100/1.30 achromat (awaiting a go for repair, succes unlikely):

"Small scratch" and extensive delamination in an Aus Jena apochromat 40/0.95 (garbage bin):

Unsetting fungal growth in an Olympus 10/0.25 achromat (repaired):

4. Using it
Counting dots/objective testing using test slides: I'm no fan:

Continues: work in progress...

"Continues: work in progress..." please continue, this is very interesting!

Have you seen http://www.microscopy.org/education/projectmicro/buying.cfm?
You can find your answer if you have such questions about buying a light microscope.

What kind of microscope should I buy? But I only have $50-100 to
spend; does that mean no microscopy? What type should I buy? What
features should I look for? I see a lot of other features advertised;
which ones are worthwhile? Can I check lens quality? What about
buying a used microscope? I see a lot of great deals on eBay. Where
do I find these microscopes?


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What’s the term used to define a terrestrial being that moves by using six limbs?
I know that Bipedal locomotion would describe how beings like humans and ostriches get around.
Quadrupedal locomotion describes how beings like dogs and cats move around.
What about insects, specifically, a cockroach? I believe a cockroach has six “legs”, what would be the term used to describe that type of locomotion?

Hexapedal. From wiki > Hexapedal

Having six legs

Note that Hexapoda is a taxonomic group including insects and the smaller groups that are Collembola, Protura, and Diplura.

Thanks. I actually searched that before the post. Based on the feedback from google, the subject must be A.) Highly complicated B.) Lacks much interest, or C.)All of the above.

What is the subject in your comment? I don't really understand your comment. Do you think 'hexapodal' is the word you were looking for?

Well, the subject is “Hexapedal Locomotion “. Not much data

Oh do you just men the term is rarely used? Yes, I am sure it is very rarely used!

I’m an electronics hobbyists, and I’m enthusiastic about robotics. Wanted to know how much man understood about hexapedal motion. Doesn’t seem like much so far.

Right.... I'm sure there's quite a bit on the subject though. But anyway.... did your question got its answer? If yes, you can click on the checkmark next to the answer.

I have seen a couple programs where professionals from universities were demonstrating big-like robots...now that I think about it, I think their “bugs” only had four legs. Hexapedal locomotion must be a complicated concept.

Searched for six legged robot on youtube and found quite a few examples. None of them are particularly fast. Anyway... talking robots is not the point of your post.


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laboratory mice, nutrition, and humans
It seems that many nutrition experiments [directed at human diet] are carried on laboratory mice. Is there any justification that nutrition that is good or bad for laboratory mice will also be same way applicable to humans ?
I wonder because mice and humans, although both being mammals, are separated probably by tens millions years of separate evolution.  By the "mammals" logic, dolphins, cows, wolves, and goats, all being mammals, shall have same diet as mice ?
Sometimes one man's food is another man's poison (lactose differences for example, due to genetic differences). That's man to man, same species. On what is "transfer of nutritional information" from species to species based ?
2) Was it ever established, or the opposite, that all humans have same best diet. Other than known lactose metabolism difference.

This is an important question I think. The commonality of genes among eukaryotes does not necessarily mean they share the same metabolism etc.

Whether using quantitative models, or "animal models", I think this is a useful quote to keep in mind:

A model is a lie that helps you see the truth.
-- Howard Skipper

As for evidence that using mice models for human nutrition is justified -- I believe there has been a good deal of research that has provided useful insight on the influence of genetics on the susceptibility to diabetes in humans using mouse models, so ... yes?

My impression is that the use of mice as human models for anything is primarily the result of historical precedent. A lot of work has been done to breed different lines of mice for particular purposes, and a lot of related methodology has therefore been established. Similarly, a lot of comparative genetic/genomic work has been done to characterize similarities between the mouse and human genomes. Developing similar methods and resources for another species could potentially require a huge investment of time and resources.
Additional reasons are purely technical and practical. Non-human primates are demonstrably more closely related to humans than mice, but breeding, storing, and working with these animals is a much more complicated issue.
And, as @SteveLianoglou mentions, any model has to be taken with a healthy level of skepticism. I prefer the quote from G.E.P. Box:

Essentially, all models are wrong, but some are useful.

No model of human nutrition is going to be perfect, whether it was modeled in mice, primates, or even other humans (as you suggested). This does not mean, however, that we cannot gain insight from the models just because they are not perfect.

+1 for the mention of practical reasons. Also, another extremely important point is that mice are pretty much the only mammal where you have the availability of an enormous amount of transgenic lines, which allow to dissect the system much more in detail. As much as I would like to work on transgenic wolves we are not quite there yet! :P

Human life span is too long and not suitable for such studies, so mice are used pretty much because their life span is much shorter and because they're already used for pretty much everything.
Mice and men as mammals have the same heritage and a metabolism which is very, very similar, almost identical - compared to the metabolism of insects, for example.
However, the millions of years of separate evolution made us the most capable omnivores on the planet, who learned to cook, became monogamous, evolved intelligence and grand-paternal care etc. We even evolved to drink milk as adults and eat gluten. So you can't possibly come up with conclusions about such complex species by looking at a simple creature that is an evolutionary dead end.
If you're using mice for your study, you can find out how something affects a mammalian species in general, e.g. radiation, but not how nutrition affects humans - because human nutrition is unique. The scientists basically don't know much about nutrition to begin with, and I suppose they're trying to figure out the basics for now, some general rules which apply to all mammals, and that's why they're using mice. A less complex organism.
But you'll get nowhere if you're researching traits unique to the human race by looking at another species which doesn't have those traits.


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Are there neurons that can sense light shining in your ears?
I know someone who bought earphones that shine light in you ears. According to what he was told, there are neurons that sense light and then make you feel wide awake when activated, which seemed like snake oil to me. Apparently the pineal gland may be able to sense light and it does secrete melatonin - a sleep regulating hormone. I'm still sceptical though as its stuck in the middle of your brain. Would shining lights in your ears be able to have any effect on how awake you feel?

This would be a great question over at Skeptics too.

Related question here but not specifially about neurons rather "extraretinal photoreception in mammals".

There is no known mechanism for light detection through the ears in humans, as far as I know. It is certainly true that the pineal gland is part of the system that regulates the circadian rhythm (briefly, the daily sleep-wake cycle). However, while the pineal gland in birds and other non-mammalian vertebrates is directly sensitive to light, the mammalian pineal gland is not (see, for review, Doyle and Menaker, 2007 and Csernus, 2006).
In all animals, the circadian rhythm is regulated by a photoperiod cue and therefore requires light detection. In mammals, the light sensors are found exclusively in the retina, the sensory portion of the eye. There are two classes of light detecting cells in the retina. First, rod and cone photoreceptors mediate vision in the usual sense of the word. These cells contain proteins called opsins that absorb photons of light and thereby excite the photoreceptors that contain them, informing the brain that light was detected.
A second class of photosenstive cells in the retina are called intrinsically photosensitive retinal ganglion cells (ipRGCs) (see Do and Yau, 2010 for review). These cells mediate "non-image-forming" vision and are an important part of the circadian rhythm pathway. They also contain an opsin called melanopsin which is a photosensitive pigment. This is not to be confused with melatonin, which is the sleep hormone released by the pineal gland. The ipRGCs in the retina send the photoperiod cue to a brain area called the suprachiasmatic nucleus (SCN). The SCN then signals to the pineal gland.
If we are generous and assume that these light-emitting headphones are the result of misunderstandings, we can guess that the confusion arises from (1) the fact that some animals have a directly photosensitive pineal gland, but not mammals and (2) that the pineal gland secretes melatonin but not the photosensitive pigment melanopsin.

Update: From a bit of research, it turns out that the company selling the headphones is not "confused" as I politely offered. I don't think this site is the appropriate forum to refute their research or claims. Suffice to say that the retina is the only part of the human brain shown to be photosensitive.

I believe there are light sensors (TRPV3) in the skin for infrared light (heat), that convey that information back to the brain from the skin.  This is kind of light detection, but it is not direct detection like the rhodopsins in the eye.
By the way, without passing information onto neurons, cells probably have a lot of sensors they may use to respond to their local environment.  This recent article talks about how olfactory receptors can be found in lung and gut cells.  so its quite possible that the conventional light detecting genes (rhodopsins) would be found in skin cells, but they may not convey information to neurons.

Can you provide a reference for a case in which a TRP channel, commonly thought of as a temperature detector, is plausibly used as a light sensor?

There has been one small study that seems to show that human skin is photosensitive.

The study I think you are referring to appeared to find that shining light on the back of the knee could entrain a human's circadian clock (Campbell and Murphy, 1998). That study has been refuted by a later study (Wright and Czeisler, 2002, see comment by Baringa, 2002). Wright and Czeisler show no response to light behind the knees. Current consensus is that mammalian photosensitivity resides only in the eyes.


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How are chromatids correctly segregated during mitosis?
In my book this is described as happening by kinetochores depolymerising from their attached microtubule spindle, then being dragged towards the spindle's centrosome by a dynein protein. I can understand this, but what it does not seem to answer is how different copies of chomatids beome attached to the correct spindles in the first place (i.e. spindles connected to opposing centrosomes). How does this work?

It's a spatial constraint.  As the DNA is replicated, the two resulting chromatids are kept stuck together by cohesin proteins.  The DNA sequence that corresponds to the centromere then coalesces the kinetochore.  It seems that the DNA-protein interactions at the centromeres creates a particular structure along the chromosome.  Since the two chromatids have their kinetochores pointing in opposite directions, when a spindle microtubule attaches to the kinetochore, it is nearly impossible for another microtubule from the same spindle to attach to the opposing kinetochore.


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

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

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

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

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

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

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

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

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

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

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

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

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

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


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When running a gel, what could cause a standard to run "faster" than usual?
We are running a gel on the products of a restriction digest to determine the size of an insert. We know the vector backbone size should be ~2.9kb; however, the standard appears to be running too quickly (or our samples too slowly), causing the backbone size to be about 4kb (which distorts our results for the insert size).
What could possibly cause this? Thank you.

Are the salt concentrations identical or similar in your sample and the standard?

Are you sure you have complete digest of the vector?

If the salt content or pH is different than "normal," run time can be affected. After all, you're applying an electric current to the sample(s) and ion concentrations and pH are sensitive to that current. Of these two, salt/ion levels are more likely to have an effect as your samples/markers are likely to be adequately buffered.


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Can I elute my GFP-tagged protein off anti-GFP antibody using a peptide?
I would like to perform a modified co-immunoprecipitation assay using a GFP-tagged protein.  We are going to bind the tagged protein to anti-GFP antibody then bind that to protein A/G beads, however we want to be able to selectively elute the tagged protein off the beads.  I know that you can elute a FLAG tagged protein off of M2 anti-FLAG antibody by adding excess FLAG, so can I use the same approach with my GFP tag?  If so, does anyone know a commercially available GFP monoclonal antibody who's epitope has been mapped so I can use excess epitope to elute my protein?  Or possibly a source of recombinant GFP?

I found a paper that describes a method to purify GFP fusion proteins using affinity chromatography: "Purification of GFP fusion proteins with high purity and yield by monoclonal antibody-coupled affinity column chromatography" (1). Unfortunately I don't have access to the full text, so I can't check how exactly they elute the GFP fusion protein from the column. They describe a specific monoclonal antibody they use, there should be information on how to obtain it in the paper.
Abstract:

GFP has often been used as a marker of gene expression, protein
  localization in living and fixed tissues as well as for protein
  targeting in intact cells and organisms. Monitoring foreign protein
  expression via GFP fusion is also very appealing for bioprocess
  applications. Many cells, including bacterial, fungal, plant, insect
  and mammalian cells, can express recombinant GFP (rGFP) efficiently.
  Several methods and procedures have been developed to purify the rGFP
  or recombinant proteins fused with GFP tag. However, most current GFP
  purification methods are limited by poor yields and low purity. In the
  current study, we developed an improved purification method, utilizing
  a FMU-GFP.5 monoclonal antibody (mAb) to GFP together with a
  mAb-coupled affinity chromatography column. The method resulted in a
  sample that was highly pure (more than 97% homogeneity) and had a
  sample yield of about 90%. Moreover, the GFP epitope permitted the
  isolation of almost all the active recombinant target proteins fused
  with GFP, directly and easily, from the crude cellular sources. Our
  data suggests this method is more efficient than any currently
  available method for purification of GFP protein.

(1) Zhuang R. et al., Purification of GFP fusion proteins with high purity and yield by monoclonal antibody-coupled affinity column chromatography. Protein Expr Purif 2008, 59(1), 138-43

In the reported paper the authors used an affinity matrix carrying a covalently coupled mAb. They then eluted the bound proteins using the standard low pH glycine buffer.


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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.


text:
Does the anterior primary ramus of T6 innvervate the Rectus abdominis?
Anatomy and Human Movement Structure and Function writes in the section about Rectus abdominis:

Nerve supply
  By the anterior primary rami of the lower six or seven thoracic
  nerves (T6/7 to T12).

What does "or" mean? Does it means that in some humans this happens and in some it doesn't?

Because muscle (mesoderm) and peripheral nerves (ectoderm) develop from different embryonic germ layers, the pattern of innervation is somewhat variable. The muscles are condensing from mesenchymal tissues more or less in place, and nerves are growing out into the muscle masses to innervate them. Because of this, the specific spinal segments that innervate muscles can vary somewhat. This is particularly true in the brachial and lumbosacral plexuses. In the brachial plexus for example:

Grayed out text for C5 in the median nerve and C7 in the ulnar nerve suggest variable contributions from those segments.
In this case, rectus abdominis is "always" innervated by at least T7 to T12. But sometimes T6 as well in some, but not all, people.

Does sometimes mean "in some people"?


text:
What is the benefit of reducing inflammation when producing antibodies?
We just learned in lecture that IL-10 promotes the formation of plasma cells over memory cells. Which seemed strange to me as IL-10 also reduces inflammation, and I figure you would want inflammation as it will help antibodies reach the site of infection.
My question is, why would the body want to reduce inflammation and produce more antibodies at the same time?

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


text:
Why is the coremata of Creatonotos gangis huge?
This Australian moth has been some kind of curse image online due to its scent organs (coremata). It is inflated to attract mates. However, why does the coremata have to be inflated for it to work, and why does it have to be so big?


text:
Trying to identify the species to which this vertebrae belongs
Found vertebrae in the "graveyard of the Atlantic" (on account of the 5,000+ shipwrecks since the 1520's) off Frisco/Hatteras, North Carolina. Tried searching online but not finding any perfect patches (although some were close with some seemingly minor differences to the untrained eye). Any ideas would be much appreciated. I have images from every angle and can take specific images if instructed on exactly what is needed. This is new for me and I am unsure who to contact if this is fruitless. If (fairly certain) identification is not possible from the image(s) please comment with any suggestions on my next contact.

I don't see any images in your question, so it is completely unclear what you are asking.

Another question about bones refers to this bone ID site which you might find helpful.

Please add more photos (one from top, bottom, front, back and each of the sides) and measurements/dimensions. Thanks.


text:
Can you identify this phyllanthus tree?
I have this tiny tree in my garden in Cancun, Q.Roo Mexico.

(More photos available here.)
The little tree started to grow in a planter. I did not pay any attention to it until it grew to 10 cm. It does grow like any bigger tree, starting with one curving branch with regular leaves. I buy unsterilized soil from locals and always get something growing without planting.
It is a phyllanthus plant, but that is all we know so far.
By now I have 20 little ones and one big one, which is a year and a half old.
It certainly is not a phyllanthus Niuri which is an incredible medicinal plant, but with way less "tree attitude" than this one.
This lovely bonsai tree could be unknown to the world, hidden in the deep vegetation of the Yucatan peninsula, one of the world's regions with the highest tree variety.
Can you identify this tree for me?

Phyllanthus caroliniensis

Phyllanthus caroliniensis, the Carolina leaf-flower, is a flowering plant native to the Americas, from the southeastern United States1 all the way to Argentina.2 The flowers are small and located where the leaf meets the stem.2

Kingdom: Plantae
Subkingdom: Tracheobionta
Superdivision: Spermatophyta
Division: Magnoliophyta
Class: Magnoliopsida – Dicotyledons
Subclass: Rosidae
Order: Euphorbiales
Family: Euphorbiaceae – Spurge family
Genus: Phyllanthus L. – leafflower

Sources:

http://plants.usda.gov/core/profile?symbol=PHCA9
https://en.wikipedia.org/wiki/Phyllanthus_caroliniensis
http://www.easttennesseewildflowers.com/gallery/index.php/Summer-Roadsides-White/Seedleaf50001
http://sura.ots.ac.cr/florula4/find_sp3.php?key_species_code=LS001041&key_kingdom=&key_phylum=&key_class=&key_order=&key_family=&key_genus=&specie_name=caroliniensis

Hello Jordi Thanks or your reply. It's close in appearance, but the stems of the P carolinensis are thicker and green. And the leaves don't seem to be  translucent as the ones from my plant.   http://i.imgur.com/U7mU1DG.jpg

the tree is identified.
http://chalk.richmond.edu/flora-kaxil-kiuic/p/phyllanthus_liebmannianus.html
Saludos
Bernhard


text:
How are nutrients absorbed by a pitcher plant?
I've read that pitcher plants thrive in nitrogen-poor soil, which is solved by dissolving insects and absorbing nitrogen that way.
I have a few plants, and when a pitcher start to rot, I cut it off and study the insects that've been trapped. I've never actually observed a dissolved insect. Both interior and exterior seems unharmed.
What digestive enzymes does the plant use?
What part of the insect gets dissolved?
What structures allows absorption and transport of nutrients into the plant? And how is it exploited?

Maybe they're just the ones that recently fell in. Older ones might have been digested. Just a guess!

No, they've not recently felled in. I usually feed them myself. Even insects placed there early in the season have not been visibly dissolved. Right now I only have one pitch left, as the plant is going into dormancy. I can post a pic of the content when the pitch start to rot.

Carnivorous plants use a variety of enzymes, including proteases which digest proteins to provide the plant with nitrogen. You might think that the hard exoskeleton would not be digested but pitcher plants (at least some types) produce chitinases which dissolve chitin (i.e. the exoskeleton), so there should be nothing left of the prey. The enzymes are secreted by tiny glands inside the pitcher.

The enzymes that have been detected in carnivorous plants include amylase, chitinase, esterase, lipase, peroxidase, phosphatase, protease, and ribonuclease.

From the Sarracenia faq.
Aside from the enzymes secreted by the plant, there are micro-organisms present in the pitcher which will dissolve organic matter.
In my experience of growing Nepenthes and Sarracenia it seems to take a long time for the prey to dissolve and there is always some material left. Which type of pitcher plant are you growing?
There's a good description of CP digestion from the ICPS.
A more technical explanation about chitinase.
I can't find any specific detail on how the plant absorbs the nutrients, though even non-carnivorous plants can absorb nutrients through the leaf surface.

Also see this SE topic on nutrient absorption via leaves


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Are there examples of the green beard effect in humans?
Green-beard effects are genes that:

Produce an effect that can be detected
Produce preference to others with the same effect

Are there any examples of this in humans?

Could you expand you question to make it more accessible to users, and thus easier to answer? What 'samples' are you interested in? Do you have an example? Thanks.

So... essentially you are looking for examples of dominant alleles with a clear phenotype? I do not understand your examples, though: racism, wars etc. are all but "easy" situations that definitely cannot be explained [just] in terms of genetics. Also, when you write "sample" do you mean "example"? I am asking because sample has a different meaning.

Essensially I am still confused. I feel that altruism toward those who are similar are very much a like altruism toward family  members. Yes, it may not be pure genetic. Then what? How, mathematically, those traits reproduce? Does every such traits come with a complementary trait to like those with the traits? Still some part is mathematically missing.

You cannot reduce everything to genetics. Ambient plays a role of paramount importance in all those situations. There are certainly a series (definitely not one) of genes that can influence how much altruist or racist or violent you are, but the education you are given and the culture in which you grow in are much more powerful in shaping your behaviour.

I've reverted all the later changes, they confused the matter more than they helped in my opinion.

Your examples are way too broad. A green-beard effect only applies to the effects of a single gene or group of linked genes, not to just any trait. Racism and your Mongol war example are too complex to be explained by this effect. In reality examples will mostly be found in single-celled organisms where expression of a single protein can have whole-organism effects such as causing aggregation, or in larger organisms where some highly visible effect is possible such as a colour change.
There are very few real known examples of the green-beard effect, but four are listed on the Wikipedia page:

The killing of red ant queens carrying a particular allele by workers with a different allele (Keller & Ross, 1998)
Altruism in social amoeba resulting from homophilic adhesion of a protein (Queller et al., 2003)
A multi-locus example in which similarly-coloured male lizards protect one another (Sinervo et al., 2006)
A neat example in yeast whereby individuals sharing an allele for a particular locus selectively aggregate, preventing those without the allele from benefiting by proxy from their trait (Smukalla et al., 2008)

References

 Keller L, Ross KG. 1998. Selfish genes: a green beard in the red fire ant. Nature 394: 573–575.
 Queller DC, Ponte E, Bozzaro S, Strassmann JE. 2003. Single-Gene Greenbeard Effects in the Social Amoeba Dictyostelium Discoideum. Science 299: 105–106.
 Sinervo B, Chaine A, Clobert J, Calsbeek R, Hazard L, Lancaster L, McAdam AG, Alonzo S, Corrigan G, Hochberg ME. 2006. Self-recognition, color signals, and cycles of greenbeard mutualism and altruism. Proc Natl Acad Sci U S A 103: 7372–7377.
 Smukalla S, Caldara M, Pochet N, Beauvais A, Guadagnini S, Yan C, Vinces MD, Jansen A, Prevost MC, Latgé J-P, et al.. 2008. FLO1 Is a Variable Green Beard Gene that Drives Biofilm-like Cooperation in Budding Yeast. Cell 135: 726–737.

Nice answer, just wanted to point out that obviously "altruism" in the amoeba cannot be extrapolated to altruism in men (not that you are saying that, but I can see such a comment coming)

@nico yes that's a good point, in this case it literally means sticking together

I asked why people are altruistic toward those who are similar to them, and they said green beard. Then here you said green beard don't work for humans.

Who is 'they'? The green beard effect simply describes one possible explanation for evolution of simple cases of altruism. It may or may not apply to humans, but no human example is known.

Perhaps you're talking about Winawer's answer here?. He doesn't mention humans, and specifically says it's more likely in bacteria, and that it probably isn't that widespread.


text:
What causes light to be brighter in the corner of the eyes?
I have noticed when in a dark room light and color seem to be brighter and more intense in the corner of my eyes. For example the light that comes from my digital clock seems like a dark pink when I look straight at it. But when I see it in the corner of my eyes it looks red.
What causes this?
Is this normal, does everyone perceive it that way?

Your retina has two kinds of light-sensing nerves: cones and rods. Cones are responsible for color vision, i.e. hue, while your rods handle differences in the value, as in how bright it is.
Humans have adapted to have cones mostly in the center of the eye (which corresponds to what you're looking directly at), where the most distinguishing color information is needed. It's not often that you'd use your peripheral vision to identify the actual color of something, and this explains why you're seeing a difference between pink and red. The intensity of the color is correct, but you aren't able to identify the hue because of a lack of cone nerves.
So yes, this is normal. A fun trick (and also totally helpful if you have a messy room) is to look a foot or so above the floor when you're walking in the dark. It allows you to see what you're walking on better, since you need intensity receptors more than color receptors.
References:
http://regentsprep.org/Regents/physics/phys09/ceyes/sensing.htm
http://hyperphysics.phy-astr.gsu.edu/hbase/vision/rodcone.html


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What is the functional difference between hemoglobin and ferritin?
I read that both hemoglobin and ferritin are for storing iron in the body. So, what is the difference between them in terms of storing/binding iron?

Why close this now? It is 5 years old and has a good answer.

Hemoglobin is the protein of erythrocytes (red blood cells) which has ferrous ions (Fe2+) bound in its subunits. These are able to keep oxygen bound which enables the cell to transport oxygen through the circulation. It's not really for storage of iron, it's for using it.
Ferritin is the actual storage protein, cells express it to store iron in case of deficiency and also to regulate the amount they have in the cell. According to this, it's mainly expressed in muscle, liver and kidney cells (but I'm not too sure about the details of that study so I might have got that wrong).
Edit: just found this in one of my old lectures; unfortunately it doesn't quote sources: ferritin stores iron in liver and heart. The total iron in the body is ~3.9g, of which 2.5g are in use in hemoglobin, 500mg in stores (an additional 250mg in the liver), 150mg in bone marrow, 300mg in myoglobin and 150mg in other enzymes. The remaining 5mg are bound to transferrin in the plasma.

yah - hemoglobin doesn't really store iron, its using the iron :)

Strange, I always thought that was myoglobin's job in muscle tissue. Anyone know myoglobin and ferritin's function in relation to one another?

According to Wikipedia we don't really know the function of myoglobin yet. Given its high relatedness to hemoglobin though, I highly doubt it serves to store iron.

@LanceLafontaine: myoglobin is essentially the muscle equivalent of haemoglobin, carrying O2 to muscle cells. Iron is used in both myoglobin and haemoglobin as the "attachment site" for O2.


text:
Macroevolution vs. microevolution
Where is the line usually drawn between macroevolution and microevolution?
I thought that, although similar processes govern both, the line was at the species level, with macroevolution being changes at the above-species level and microevolution being changes within a species (something like changes in allele frequencies).  (So it's a question of timescales?)
However, I heard someone talk about microevolution among different species of Drosophila, specifically referring to copy number variation.  Was the choice of "microevolution" to describe this process (more likely) due to the similarity between Drosophila species or the type of change in the gene? Or at this level, is the distinction rather arbitrary?

Let's start with a word of caution: on the internet, the terms macroevolution and microevolution (especially together) are usually used primarily in creationist rhetoric. As such, it is usually best to avoid them, especially when talking to a lay audience. The main mistake creationists perpetuate when thinking about micro-vs-macro evolution, is that the two are somehow different and distinct physical processes. This is simply not the case, they are both just evolution. The scientific distinction between the terms, comes not from the physical world around us, but from how we choose to talk about it. When a biologist says "microevolution" or "macroevolution" they are actually signaling what kind of questions they are interested in asking, or what sort of tools they plan on using.
Verbal and empirical theories
In verbal and empirical theories, the micro-macro distinction is usually one of timescales. A person in the macroevolutionary paradigm, usually asks questions above the level of individual species, as Evolution 101 writes (emphasis mine):

instead of focusing on an individual beetle species, a macroevolutionary lens might require that we zoom out on the tree of life, to assess the diversity of the entire beetle clade and its position on the tree.

Empirically, macroevolutionary answers to these sort of questions are usually ones that don't have access to detailed evolutionary histories or direct experiment. Instead, the method tends to be ones that use geology, fossils, and back-inferences from broad differences/similarities of existing species. As such, most macroevolutionary theories tend to be descriptive, instead of predictive. Most of paleontology can be classified under the macroevolutionary paradigm.
If someone explicitly says that they are looking at microevolution then this usually refers to a contrasting methodology that tends to be heavy on direct experimental manipulation. Most importantly, microevolutionists tend to have access to rich and detailed evolutionary histories. As such, it is not surprising that you saw a study that identified itself as microevolutionary while looking at different species of Drosophila. The fruit fly is one of the widest used model organisms, and as such the study you are referring to probably experimentally manipulated populations of fruit flies and collected a rich dataset of genetic changes in the generations of fruit flies that they studied.
Of course, most studies are at intermittent levels, and no this isn't called meso-evolution (except by silly people). If you are not clearly using the macroevolutionary nor the microevolutionary paradigm but still looking at evolution, you would just simply say 'evolutionary' without any prefix.
Formal and mathematical models
For formal and mathematical theories, the micro/macro distinction is also one of methods used by the theorist not of the underlying domain they are studying. Mathematical modeling of evolution falls into two broad categories: frequency-dependent and frequency-independent models. Frequency-dependent models almost never make the micro-macro distinction. Although you could argue that dynamic models in evolutionary game theory are micro, and static equilibrium concepts like ESS are macro, but I doubt theorists would endorse this view. As such, I will focus on the frequency-independent models.
For frequency-independent models, they key concept is the fitness landscape -- a way to map each genotype to a fitness. If the model tracks a population (a distribution over vertices in the fitness graph) on a static fitness landscape, then it would typically be just called an evolutionary model (the word microevolutionary is seldom used explicitly in this field). However, if the authors assume that mutations are extremely rare and thus any mutant goes to fixation before a new one arises then they can use Gillespie (1983, 1984) to replace the population by a single "typical individual" occupying one vertex of the fitness graph. At this point, the model becomes macroevolutionary and the underlying rule for calculating fixation probabilities would be the microevolutionary component. This approach is often also coupled with an alternation of change in fitness landscape followed by mutations and a selective sweep. Fundamentally, though, macroevolutionary models are just convenient (or tractable) approximations to a real underlying evolutionary dynamics. This should never be forgotten.
Strange example bridging the gaps
Finally, it is important to stress that the macro- and micro-evolutionary paradigms are not necessarily exclusive and do not have to correspond to a difference in timescales! This is best done with an example of a respected theoretical study that mixes everything together. Kauffman & Weinberger (1989) used their newly developed NK model or rugged fitness landscapes to study maturation of the immune response (Tonegawa, 1983). The developed a macroevolutionary mathematical model because they used Gillespie's trick to replace a population by a typical individual by abstracting away from the underlying microevolutionary calculation of fixation probabilities. However, their model was studying evolutionary dynamics within the human immune system (so timescales of days to weeks) and was tuned by parameters gathered by empirical microevolutionary studies that tracked individual nucleotide changes (Crews et al., 1981; Tonegawa, 1983; Clark et al., 1985). Lastly, the study results can be used to inform a question typical of verbal macroevolutionary theory: Are there any examples of sudden leaps in evolution?.
As such, the above study used a formal macroevolutionary model, informed by empirical microevolutionary work, to help us understand a question typical of verbal macroevolution while looking at a physical process that operated on the incredibly short timescale of days to weeks. No wonder people are so confused by the micro-macro "divide"!

References
Clark, S.H., Huppi, K., Ruezinsky, D., Staudt, L., Gerhard, W., & Weigert, M. (1985). Inter- and intraclonal diversity in the antibody response to influenza hemagglutin. J. Exp. Med. 161, 687.
Crews, S., Griffin, J., Huang, H., Calame, K., & Hood, L. (1981). A single V gene segment encodes the immune response to phosphorylcholine: somatic mutation is correlated with the class of the antibody. Cell 25, 59.
Gillespie, J.H. (1983). A simple stochastic gene substitution model. Theor. Pop. Biol. 23, 202.
Gillespie, J.H. (1984). Molecular evolution over the mutational landscape. Evolution 38, 1116.
Kauffman, S. and Weinberger, E. (1989) The NK Model of rugged fitness landscapes and its application to the maturation of the immune response. Journal of Theoretical Biology, 141(2): 211-245
Tonegawa, S. (1983). Somatic generation of antibody diversity. Nature 302, 575.

Thank you for the well-referenced and very detailed answer.  I did try to look this up myself but kept getting creationist websites, so I was hoping you might answer :)

@dd3 no problem! It was a fun question and got me to think more carefully about the micro/macro distinction and actually write something down. I might end up making a blog post out of this later, so thanks for that! :D