text:
RNA vaccination and Autoimmune Reactions
To my knowledge we do not have any robust experience with RNA vaccination. Most likely this will change in the near future as RNA vaccines against COVID are in the pipeline. The rationale behind this method: Viral mRNA is injected (I guess in a muscle), the mRNA is translated (in muscle cells, for instance), the protein is exposed on the cell surface to cause a response of the immune system. As we know proteins tend to interact and build complexes. I speculate that this could easily lead to an immune response against cellular protein structures.
Is there any knowledge or speculation concerning an autoimmune disease in succession of RNA vaccination?

I would imagine the proteins are secreted instead of being cell-bound, just for this very reason.

Welcome to SE Biology. I think it is important to realize certain things about this site. First, this is a question and answer site, not a discussion site — answers that are mere speculation or opinions are generally off-limits. Second it is a site concerned with specific aspects of Biology (see the [Tour]) rather than medicine — for which there is a sister site SE Medical Sciences. In my opinion your question should have been submitted there, if at all. I say, if at all, as there are recent authoritative articles which I would try first. E.g. Go to nature.com and search for covid vaccine.


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What are the advantages of mRNA vaccines?
When the mRNA vaccines for COVID-19 are administered, mRNA molecules are introduced into the cells of the subject. The translation of this mRNA determines the productions of antigens, which in turn stimulate the immune response.
Wouldn't it be simpler to have this translation happen in vitro, and then inject the antigen into the subject's body? What are the advantages of making the translation happen in vivo instead?

Quick answer - typical protein-based subunit vaccines tend to produce a strong antibody response (humoral immunity), but not a very strong cellular response. By using an mRNA vaccine that produces the viral protein of interest (in this case the SARS-CoV-2 spike protein) in cells, a strong cellular response can be elicited in addition to the humoral response. This allows for the body's full defenses to be used against an infection right away. I don't have time to give an expanded answer right now, but I'll try to later.

see paragraph here: https://en.wikipedia.org/wiki/RNA_vaccine#:~:text=The%20benefit%20of%20using%20mRNA,COVID%2D19%20in%202%20days.

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text:
How could I make my fish gold?
I'm doing research on how the pigmentation of fish. I want to know how could you change it with the help of genetic modification. I have some Silver Dollar fish (Metynnis argenteus) and I want to make them have a shimmering gold appearance like a gold piranha. I was told by someone on reddit that it would be much easier to breed them to create the desired phenotype, but I don't think that's possible since they don't produce many observable differences in pigmentation. So, I want to know if anyone has any experience with this and could help me out. I'm still trying to understand all of this so it would help to not be too technical.

Did you figure out how the gold appearance in a gold piranha happens? Is it a certain chemical or rather a physical effect?

I think you need to clarify what kind of research you are doing. Breeding and genetic modification might both produce the desired result, but in research it is likely not the main point (but instead getting to this result in a certain way and obtaining the insights from doing this).
There are many studies on coloring of zebrafish and guppies, producing all kinds of differently colored mutants... but I doubt that one can predict the result in advance, since the link between the genotype and the phenotype is not well understood yet. In fact, understanding this link is precisely the point of these studies. You can start with exploring the work done in this domain by Nüsseln-Volhardt's group, see, e.g., here.

Im trying to find information on what genes cause the coloration in charcins. I also want to know if all fish have the same genes for pigmention. I know that the relation between genotypes and phenotypes are not fully understood yet. I'll try to find out if guppies and danios (zebrafish) share the same genes. If they do there is a possibility that piranhas and silver dollars do as well.

@PrinceZay You'll get the best answers here if you ask the questions that get exactly to the answer you want. If you want to know "if all fish have the same genes for pigmentation" then do some research on pigmentation and ask about it. Asking "how can I make my fish gold" won't answer the other question. We call this the XY problem at StackExchange.

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Can forensic DNA analysis be used to generate a visual approximation of a suspect?
In light of the current US supreme court case, I'm curious if enough information can be teased out of a DNA sample to get a "reasonable" approximation of the suspect (never mind the legality). I realize the term reasonable is subjective, so characteristics, such as skin color, hair color/texture, approx height, eye color, freckles, etc. I came across this article a few years ago, but the company that made the test went bust in 2009. It could detect race (how accurately?). I seem to recall reading an article a few years ago stating that "soon" we'd be able to get an approximation of appearance from a DNA sample that was better than a sketch artist could provide, though I can't recall now where I read it.

In theory it's possible to have an approximation, but not to know with certainty. Identical twins have the same genomes and look very much alike. Whether it can be done in practice depends on how well we can model the relationship between genes and looks and on how much information is necessary for a judge to permit arresting and questioning a suspect.
Scientists know relatively little about how genes influence physical appearance yet. The height of the suspect is controlled by hundreds of genes with complicated interplay between them, so I don't think it's possible to estimate it just yet. Eye color is more simple: it's controlled by only 3 genes. We know weight is partially controlled by genetics but exactly which genes are responsible is not clear. Age is impossible to tell by DNA alone. Skin color should be predictable, since it's caused by melanin production, and we know which genes are responsible.
So to sum up, DNA analysis can tell the gender, skin color, and eye color of the suspect, but very little about height and weight, and virtually nothing about age. That's not enough information to identify a suspect, although perhaps someone with legal knowledge could comment on this.
But let's say in the future we know enough to determine all these traits. Then DNA analysis would be helpful but never enough to identify a suspect. The problem is that different cells in your body undergo different mutations in their DNA with time. That's how one gets cancer. Let's say there's a mutation in the eye color gene in skill cells from a murderer found on a victim's body. It won't affect the murderer's eye color but it will affect the forensic analysis. So it's impossible to know for sure what the suspect's appearance will be.

I agree with most of your assessment. I'd think that some genetic markers could give an approximation on traits like height though. Clearly diet makes a difference too, but working under certain assumptions I'd think a rough estimate could be made. Fair point about mutations, but I'd be curious to see just how significant those turn out to be on the whole. There would clearly need to be all kinds of other traits analyzed as well. Things like cheek structure, lips, nose, hair texture, freckles, widow's peak, etc, etc. Some of these would be easier to glean than others...

@single_digit You're right to point out that one can make a rough estimate, and perhaps it will get more accurate in the future. One of my main points was actually that a judge would not consider the information enough at present to issue a search or arrest warrant, for example, so it's unlikely that these methods would help in an investigation. I have no formal legal knowledge or experience, though. I'm curious for feedback on this from someone who knows more about the law.

For readers coming along later, it appears the remarks in this answer are now out of date with respect to determining age from DNA.  While not a precise measure of chronological age of the individual from whom the DNA same, it appears a Horvath Clock or other epigenetic clocks can actually give reasonable results in many cases. See this Wikipedia summary as one starting point:  https://en.wikipedia.org/wiki/Epigenetic_clock

As far as I know there are ongoing efforts to find genes that affect forensically relevant traits e.g. facial characteristics and fingerprint pattern type/ridge count, and it's definitely a topic of interest to some law enforcement agencies. However as previously mentioned, there are many factors that could influence complex traits, including in utero environments and other, postnatal effects, such as nutrition, as well as the interaction between those environments and genetic influences. Given past literature showing jury-eligible members tend to over-estimate the importance and statistical accuracy of DNA profiling, however, I think even with future technological and methodological advances in genetic research, this type of evidence should be used with caution, explained with extreme care and thoroughness, and never taken as the single determining factor in reaching a verdict.

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Do DNA supply houses maintain a watch list for dangerous oligos (Anthrax, Smallpox, etc.)?
The recent work by DeLoache, et al. on a synthetic opiate-precursor production pathway in yeast has generated a lot of scare stories in the media about people homebrewing heroin as easily as they would beer.
In connection with DeLoache and coworkers' research, the New York Times has claimed

Their DNA could be put on a watch list, as sequences for anthrax and smallpox are, so any attempt to buy them from DNA supply houses would raise flags.

I have never heard of such a watch list, and a thorough googling turned up nothing relevant save for the NYT article itself. Does this watch list exist?

I have heard about this before. But this is kind of pointless when you know how to handle and manipulate DNA. Besides that, it is not trivial to puzzle a virus together.

@chris But this is kind of pointless when you know how to handle and manipulate DNA. Yeah, that's my real question: how could such a list even work? How would it handle silent mutations? How would it handle SNPs, or other mutations easily fixed with PCR? Combinations of SNPs?

I think it will hold off people with no real knowledge. It won'T be a big hurdle for everybody else.

Cross-posted to Skeptics. In addition, while the smallpox genome might be within reach of gene synthesis, having an anthrax genome does not bring you any closer to synthesising an anthrax bacterium.

i encountered same question. nowadays you can order very big piece of dna custom-made. some viruses are <10Kb so they are easily synthesized. anthrax is 5Mb and smallpox is around 180Kb, so those are hard to produce in vitro

I guess the fear is not necessarily to have someone produce a viable virulent anthrax from a DNA sample, but to have them introduce virulence factors of those watched pathogens into previously harmless organisms they could easily buy and grow.

@aandreev While the whole bacterial genomes are far larger than a typical gene synthesizer can sell, DNA for the dangerous parts, like the botulism toxin protein, are small enough to be made. These could potentially be inserted into some other bacteria, like E. coli pretty easily. If you can do a little more work, you might be able to get into a cold or flu virus.

@aandreev Additionally, positive strand single stranded RNA viruses have genomic RNA that works as mRNA. If you had the DNA for the virus, and made the RNA through in vitro transcription, that RNA would be enough to infect cells and make virus, assuming you could get it into cells, which is still pretty hard. But maybe the RNA could be produced and stored more easily than the whole virus.

it seems to me that problem of producing virulent/pathogenic (i might mix terminology here) agents from scratch (via ordering DNA pieces and cloning) is medium to hard, but definitely doable. question is does companies check sequences and report to FBI

@aandreev I think they do. You could always get the smallpox sequence and submit it to genscript for synthesis. If the FBI comes knocking on your door, let us know before they send you to gitmo.

@user137 sure thing

I don't know how generally such things are followed, but in at least one contract gene synthesis provider's FAQ, they address this issue, and mention two such government-provided resources:

Q: Does GENEWIZ have a procedure in place to address potential biosecurity
          concerns for gene synthesis projects?
GENEWIZ is fully aware of all risks and benefits associated with gene synthesis research. With the knowledge that gene synthesis technologies have the ability to enable de novo reconstruction of dangerous pathogens, GENEWIZ employs rigorous quality control policies and procedures to safeguard against abuse of the genes we synthesize. GENEWIZ actively monitors the Screening Framework Guidance for Synthetic Double-Stranded DNA Providers [link], drafted by U.S. Department of Health and Human Services (HHS), as well as the Select Agents and Toxins list put forth by the U.S. Government.

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Why are Mendelian mutations considered rare?
I was reading the Wikipedia articles about genome-wide association studies (GWAS) and whole exome sequencing.
As I understand, GWAS is used to identify most common SNPs that are proved to be associated with a disease/disorder. On the other hand there is an article about whole exome sequencing method where it said that it is more sensitive to identify Mendelian mutations in contrast with the SNP genotyping methods used in GWAS.
As the graph shows below, there is a distinction between Mendelian mutations and common mutations. What exactly does Mendelian mutation mean (why do we call it Mendelian)? What makes a Mendelian mutation so rare but effective in contrast with common mutations? Aren't common mutations inherited with the same rules as Mendelian ones?

I will focus more on the question of rarity, which the comments have not addressed. Recall that natural selection is operating on the human population. Because of this, bad mutations of large effect (disease mutations such as CFTR in your figure) are very rare, because selection is removing them at some rate from the population (people with cystic fibrosis are unfortunately very sick and are not as fertile as healthy people), whereas mutations with very little effect are not subject to much selection. Paper one and paper two are reviews describing the population genetics in more detail.
As suggested by commenters, Mendelian mutations are so named because they are single loci of large effect controlling traits (approximately). Common variants, by this logic, must have very little phenotypic effect, because they are not purged by natural selection (assuming that they are deleterious, or disease-causing).
Rule of thumb: 'Mendelian' alleles can easily be used to predict phenotypes (like disease) and are acted upon by natural selection. 'Common' alleles are individually not informative about phenotype and are thus mostly ignored by natural selection. Not exactly true, but close.

In reality, Mendelian variants are very rare compared to non-Mendelian variants. Why would evolution evolve dominance or recessiveness? If you didn't learn about this in grade school, it would likely come off as a very weird and specific thing to evolve. More likely, there is an optimal value for whatever phenotype you are going for (enzymatic efficiency, height, disease, etc.), and it wouldn't make sense to want 25% of the population to have the trait completely turned off and the other 75% to have the trait completely turned on.
Mendelian alleles only account for a fraction of the variance in most diseases, because they are so rare, whereas common alleles, taken together, explain the bulk of the genetic variation in most diseases. However, when there is a Mendelian variant, its effect is so large that it is obvious. This makes them easy to study.

I'd like to see some sources for your answer, specifically your claim that Mendelian variants are very rare.

@bob1 There are certainly much fewer than 100,000 Mendelian variants. Even if we say there are only 10 million SNPs (some sources say 600 million), Mendelian variants are 100x rarer.

"Why would evolution evolve dominance or recessiveness?" - recessiveness basically evolves from regulation that is useful whether or not a defective copy of a gene is present. Typically recessive alleles are recessive because they cause a defect in function and because the transcription/translation of the other copy happens to a level that doesn't depend strongly on the number of copies. Dominance similarly doesn't evolve directly, it's just a consequence of a gene product having an effect.

I think that "Mendelian" is being used by the OP due to confusion, not because they are interested in the population dynamics of dominance. I think that what you are talking about may be true in the context of the genetic architecture of traits and we have to some extent discussed that here. But they seem to be more interested in the dynamics that lead to the plot that they show, which is basically the result of selection.

@MaximilianPress Yes, absolutely. I meant this more like a footnote to your answer

@BigMistake I think it would be appropriate to say that there are <100,000 Mendelian variants that we know of. I really really doubt that we have studied enough genomes to know whether many of those SNPs is Mendelian or not, even with GWA studies going on.


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changing DNA of an organism
is it possible at least theoretically to change the DNA of an organism ?
so we take for example an organism with multiple cells then we change its DNA without killing this organism
to correct genetic defects or to improve it

See this.

Yes, it is possible, but you have to engineer the totipotent stem cell that forms all the other cells in the organism's body. There is no way to alter the DNA of every single cell in a human or mouse, for example, after they have begun developing.

Erm. Sort of. Kind of depends on your definition. It hasn't been done for multicellular organisms though.
Craig Venter et al did this exact experiment a few years ago with a completely synthetic Mycoplasma genome.
http://www.jcvi.org/cms/press/press-releases/full-text/article/first-minimal-synthetic-bacterial-cell-designed-and-constructed-by-scientists-at-venter-institute-an/
If you just want to change small segments of DNA that's where cloning and gene therapy come in, and we do that A LOT.


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How to prepare sample for SDS PAGE?
I usually take my protein sample 0.8ml and sample buffer(2X) 0.2ml for my sample preparation in SDS PAGE. Am I using correct proportion? my protein sample concentration is 4.1mg/ml. What is the standard protocol?

Why do you dilute like this? If you do so, the final concentration of your sample buffer is 0.5x. For this dilution you need a 5x sample buffer.

@Chris. Yes, my final concentration become 0.5x. Is it effect on getting bands on SDS PAGE?

Surely it will affect the gel - the concentration of the ions is lower than it should be, so your gel can run funny.

@chris, Can you please give me protocol for preparing sample buffer? Because in different protocols showing different composition.

Sample preparation for protein gels is not a complex task. Simply mix the appropriate amount of sample buffer with your sample and load it. For a 2x sample buffer use equal amounts of sample and buffer, for 5x sample buffer use 4 parts of sample and 1 part of buffer (for examle 40µl + 10µl). Heat the mixed samples for 5 minutes at 95°C, cool them immediately on ice and load an appropriate amount of it on the gel. How much is possible really depends on the type of gel you use - 20-30µl should usually be possible.
Do not load too much protein, otherwise you will not be able to seperate them appropriately. I usually use between 20-30µg of total protein per well or as little as 50ng for purified proteins (this depends on the antibodies as well).
Have a look on this webpage for some further information.
For sample buffers I usually use the following recipes. I prefer the 5x, because I can load more protein, if the preparation is not very good. I also prefer the use of DTT to Mercaptoethanol, as it is much less smelly. If you have no DTT (or prefer mercaptoethanol) it can be replaced by an equal amount (200mM in the 2x, 500mM in the 5x buffer). If you use DTT, you need to make aliquots of the buffer and store it at -20°C, a working aliquot can be kept at 4°C for two weeks or so.
2x sample buffer:

4% SDS
20% glycerol
100 mM Tris-Cl, pH 6.8
2 mM EDTA (ethylene diamine tetraacetic acid)
200 mM DTT (dithiothreitol)
0.1 % (w/v) bromphenol blue dye

5x sample buffer

10% SDS
50% glycerol
250 mM Tris-Cl, pH 6.8
5 mM EDTA (ethylene diamine tetraacetic acid)
200 mM DTT (dithiothreitol)
0.25 % (w/v) bromphenol blue dye


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What is SDS PAGE gel polymerization time?
I am working on 20% SDS PAGE. I want to know optimum polymerization time for 20% resolving gel and 6% stacking gel. If I increase the time then would it affect the band pattern?

it depends on many factors such as APS concentration, TEMED concentration and ambient temperature

APS and TEMED concentrations matter a lot. Even with that standardized, your polyacrylamide on the shelf is losing reactive groups as we speak. Who know how long it sat in the warehouse? It all adds up to the fact that the minimum required time is practically impossible to predict.
On the other hand, once polymerized, a gel is stable for days without apparent loss of quality. Overnight polymerization will be "optimal" in the sense that the gel won't get any better with longer or shorter polymerization. But so will be a two-days old gel. People used to cast gels every Friday, and leave them in wet paper towels in the cold room for the next week.

There is nothing like a general polymerization time, as this is a chemical reaction which depends on many factors. The most important are:
APS: Concentration and and the age of the chemical are important. APS decomposes over time when it is not stored completely dry (which it is not on most benches) and also with freezing thawing cycles in storage. APS provides the free radicals necessary for the reaction; without it no polymerization will happen. In my experience APS is the most critical part of the reaction, with old solutions making the polymerization really slow. I would make a stock solution, freeze it in aliquots. For making gels I thaw on aliquot and keep it in the fridge afterwards for a week and then discard it.
TEMED: Is the catalyst for the reaction, too much of it will start the reaction very fast.
Temperature: The warmer it is, the faster the polymerization will be. In the cold room you will not get complete reaction, if any.
What you can do is to store the remaining acrylamide in the beaker and wait until it is polymerized. Then your gel should also be ok to use. A longer polymerization time (assuming that your gel was completely polymerized before) will not change the resolving capacity of your gel.

"with old solutions making the polymerization really fast"... I guess you meant fresh solutions. Old solutions are bad.

-1 For having to scrape acrylamide out of the beaker! (I didn't actually down vote)

@canadianer You can do the mixing in a plastic tube if you want and discard it when the polymerization is finished. Makes more trash though.

Others have already explained the effect of different factors on polymerization time. This answer is mostly about:

If I increase the time then would it affect the band pattern?

It may.
Some tips:

Make a little extra mix so that you can leave some in the beaker to know if polymerization has happened.
Always layer the poured resolving gel with water so that the gel doesn't get dried up.
Don't leave the gel for long time after polymerization. Not only it wastes time, it also may cause the gel to dry up and SDS/Urea to precipitate/crystallize.
Wash the gels with the running buffer after removing the comb/before loading the sample. You can use a fine needled (high gauge) to do that.
Pre-run the gel to equilibriate it; this not generally necessary and as Nick Sandor said, would destroy stacking gel. However it is important for big urea gels and somewhat important for continuous SDS-gels if you have left it for a quite some time post polymerization. So it is better to not leave the gel for a long time

In theory, pre-running SDS-PAGE discontinuous gels will destroy the stacking gel, because glycine, which doesn't belong there, will go in. In practice, I see a lot of people recommending it, regardless of theory, and in the absence of any empirical evidence. "To remove unpolymerized acrylamide", some say, to which I reply that it will take just as much as passing bromophenol blue and glycine through the gel.

@NickSandor For usual SDS-PAGE it is not necessary- I agree and thanks for the point; it would change the pH and composition of stacking gel. I edited the answer. But for urea gels (continuous) it is always recommended.

The polymerization time is strongly dependent on APS and TEMED concentrations. I do not think that the increased time for polymerization would somehow influence the division of the proteins in negative way. Actually the time varies from 20 min to 1 h, but I personally leave the gel to polymerize longer. Unpolymerized Acrylamide could build adducts with some proteins which could result in multiple bands. In addition acrymamide itself is toxic, so I always give it additional time to polymerize. Good luck!


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can a bacterial protease inhibitor cocktail be used for Western Blotting involving HUVEC cells and HT-29 adenocarcinoma cells?
Dear fellow biochemists,
I need some advice on Western Blotting, more specifically the use of certain protease inhibitors with the RIPA cell lysis buffer and protease inhibitor cocktail. A Millipore technical representative recommended my using "539134 | Protease Inhibitor Cocktail Set III, EDTA-Free - Calbiochem" as a good general protease inhibitor cocktail.
I have another protease inhibitor cocktail which includes: AEBSF, bestatin, E-64, and pepstatin A (E.coli protease inhibitor cocktail) but it does not include the aprotinin or the leupeptin inhibitors. Is there a risk of proteolysis in a mammalian cell lysate without these latter 2 inhibitors? The specific cell types which we are using are: HUVEC (human umbilical vein endothelial cells) and HT-29 adenocarcinoma cell line.


text:
Will gene mutations give rise to a protein that is shorter than the full length protein?
What is the relationship between deletion or insertion of nucleotide in the gene sequence and the protein's length that is coded out from the mutated gene?

It depends on the nature of the insertion or deletion.
Many mutations are point mutations, where one single nucleotide is changed:
ATGACTACGCTATTCAGT
M  T  T  L  F  S

ATGTCTACGCTATTCAGT
M  S  T  L  F  S

Notice that changing the fourth A to a T caused a T->S change in the final protein, and didn't change its length. However, an insertion/deletion is different. Starting with this sequence:
ATGTCTATTGACGCTATTCAG
M  S  I  D  A  I  Q

Let's have an insertion, an A after the A in position 7:
ATGTCTAATTGACGCTATTCAG
M  S  N  STOP

Notice we now have a stop codon (TGA). This means the protein will be truncated, and will be much smaller. That isn't always the case. Rather than adding an A at position 7, let's add it after position 9:
ATGTCTATTAGACGCTATTCA
M  S  I  R  R  Y  S

The protein is about the same length, but the AAs are completely different than the original version after the insertion:
M  S  I  D  A  I  Q
M  S  I  R  R  Y  S

This is called a frameshift mutation. Biologically, these mutations are harmful because they change all the amino acids after the mutation, the protein is often not functional and can end up being different sizes (if a stop codon is now in frame or out of frame) and the protein can even end up being unstable and rapidly degraded.
So, to answer your question about the length of the protein, it'll increase/decrease by one AA every time 3 nucleotides are added/removed, but can increase or decrease drastically if a stop codon is added or removed by a frameshift mutation.


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Do drugs' levo isomers have a better interaction with the receptors in our body than dextro isomers?
Examples of levo drugs include levothyroxine, levocitrizine, and levodopa. Is there any specific reason why the receptors in our body exhibit this stereoisomerism and hold a high preference for the levo isomer form of any drugs?

I can't answer this definitively, but I believe it's because the amino acids that make up the normal binding partner for the receptors are levo.  So the parts fit together better and easier than trying to me dextro versions.  It would be interesting to know if the HTS and Fragment libraries used in modern drug discovery even include D- rotated molecules for this reason.

This is purely coincidental. The term levo simply means the direction that the pure enantiomer of the compound rotates plane polarized light at a specific wavelength and has no direct bearing on the interactions with biological systems. A quick search of the Dictionary of Drugs database gave 6845 compounds with optical rotation >0 (dextro-compounds) and 8406 compounds with optical rotation <0 (levo-compounds). So there are plenty of chiral drugs that rotate plane polarized light each direction.
The inclusion of 'levo' (or 'dextro' for that matter) into the name of the compound was done to distinguish enantiomers of chiral compounds. This naming convention as fallen out of favor due to the improved ability of chemists to know the absolute stereochemistry (e.g., R/S) and distinguish enantiomers based on their structure.
Dextroamphetamine (Adderall) and dextromethorphan (Robitussin) are a couple of examples of drugs with 'dextro' in their names.
With respect to the stereochemical nature of HTS and fragment libraries, most compounds are achiral (do not rotate plane polarized light). I'm not aware of any library biased toward optical rotation, and I'm not aware of any chemist making any selection of compound based solely on an optical rotation measurement. In fact, chiral compounds are often studied first as the racemate. If there is activity, the individual enantiomers are tested.


text:
Problem with bacterial transformation with electroporation
I have a problem with a bacterial transformation of a yeast gene that I can not solve.
I isolated yeast DNA and did a PCR to get my product. I am using pCGCUm vector with a GFP construct. I digest both my plasmid and my product with SmaI and ClaI for 4h each. After that I purify the product from a gel and perform a ligation with high efficiency T4 ligase for 30 minutes at 22°C. Then I transform my electrocompetent cells with electroporation (2000V, 5ms), recover it in 0.5ml LB for 2h and plate it on LB-A plates.
First time I only did:

insert1
insert2
Empty vector

and it did not work (empty plates for insert and confluent growth (after 18h) of some weird colonies on EV).
I did the same ligation again and this time I included controls
This is what I did:

insert1
insert2
Empty vector
negative control (EK cells + electroporation)
positive control (EK cells + undigested vector)

Now I got:

insert1 (no colonies)
insert2 (big colonies a lot of them)
Empty vector (big colonies a lot of them)
negative control (EK cells + electroporation) (no colonies)
positive control (EK cells + undigested vector) (big colonies a lot
of them)

The negative control suggests that my competent cells are ok, at least they are not resistant to AMP.
What confuses me is that even with positive control I get some weird big colonies - if I got nice transformants here I would assume that either digestion of ligation was not successful, but now I doubt that).
Another thing that is weird is that now I got that colonies also on my insert2 plate.
The scan image of big colonies looks like that:

I have a question if anyone has had that kind of colonies? I was thinking of maybe having contaminated cuvettes for electroporation but I had them in 100% EtOH for 3 days and then dried in fume hood for 3 hours before electroporation.

Some questions: 1) Can you describe the pCGCUm plasmid in more detail? antibiotics, presence of ccdB gene or MCS, etc. Can't find it on the web. 2) What are EK cells? 3) Where did you get your transformation protocol? From someone else in your lab who's had it work for them? I've been told not to let cells recover from electroporation for more than 1 hr because they may lose the plasmid during prolonged recovery, but if this has worked for a labmate then that's not an issue. 4) Have you repeated the 2nd expt (with the extra controls)? If it's contamination it might not reappear a second time.

I agree those big colonies do not look normal. It's possible that your competent cells are contaminated with some strange strain. You could try plating serial dilutions of your competent cells (at least 10^6 fold dilution) until you can see individual colonies, and see if it's a mix of normal looking and weird colonies. Again, ask other people in the lab if they've had contamination issues--one time I struggled for a long time with low tranformation yields until I learned someone else was having the same problem, and we made a new batch of competent cells and were fine.

Hey. Thanks for comments. pCGCum is 5.5kbp long. It has an ampicillin resistance and MCS long 53 nucleotides - SmaI on 8 and ClaI on 47. EK cells are electro competent cells(sorry I translated it from german). The protocol is standard protocol that has been used in our lab for long time now and has worked ok till problems appeared lately. At the moment it is only 2 people that were actually doing it and other one have the same problems with that colonies.

We made a new batch of electro competent cells and alkaline phosphatase is new. Ligation kit was changed before my experiments. I will make new batch of chemical competent cells and try with those and hope to have some success. I will also do plasmid prep on insert2 plate where I have the colonies and try to digest it, maybe they are just "weird positives" - which would still not give me the answer but at least I will know something more. I will also plate different dilutions next time repeating exp.2 with our EKcells, EKcells from other lab and chemical competent cells.

I did try another transformation:

again with our electrocompetent cells
Cells form other lab
chemically competent cells I made

The good thing is that on chemically competent cells there is no weir colonies, however efficiency of transformation was very low, but still, I think it is ok - will confirm it in next day if I really got the right insert.
On the other hand my electroporation still did not clear out. I got colonies on both our cells and cells from other lab, however in much lower amount there. I will try to change the cuvettes and try to use different electroporator as this seems to be the only thing that could make problems - although I am still not convinced about that :)
Thank you for your help and I will report the progress so if somebody else will have that problem again he could maybe benefit from my findings.

So, the answer is clear now. The contamination was in cuvettes. Some kind of bacteria that can survive 70 and 96% Ethanol, probably in form of spores. What I did is I putted cuvettes in LB medium for several hours so I got growing cells, than soaked them in 70% ethanol overnight and rinsed them with 96% EtOH for 2 hours. Than I added LB to cuvettes, "recovered it on 37 for 2 h and plated on LB plate. Plate was clean next day and also I did the normal transformation which worked ok!

Do you re-use cuvettes? If you do so, you should at least incubate and wash them in 0.1M HCl to get rid of DNA contaminants before washing them with 1% SDS and getting rid of all the detergent afterwards with a lot of water. We stopped doing this and either use the cuvettes only once when we really have to do electroporesis and the rest of the time chemical competent cells work fine.

Yes, we reuse the cuvettes and we only washed them with ethanol. Everybody in the lab were doing it that way, but with all the problems I had with cloning I will use chemically competent cells from now on. The efficiency is the same and I like it more.


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Do individual (free) nucleotides base-pair with other free nucleotides?
I had a student ask me about this and my google-fu let me down. He asked if individual nucleotides (not in a nucleic acid) base-pair with their complementary nucleotides, essentially forming many dinucleotides. I would think this would occur, but have no evidence one way or the other. If it does, how does this impact the usage of nucleotides in nucleic acid synthesis? I'd assume they have to be broken apart first...

The base-pairing in solution only provides 2-3 hydrogen bonds, which the nucleotides could also form with water itself. There are no stacking interactions between bases in water like in a DNA helix. So I strongly doubt that the nucleotides would form a significant amount of dimers in water.
I found a paper confirming my assumption:

Short nucleotide fragments such as mono- and dinucleotides are
generally unable to form stable hydrogen-bonded base pairs or duplexes
in water.
Minimal nucleotide duplex formation in water through enclathration in self-assembled hosts.

An easy way to confirm this would be to observe the proton NMR spectra of free nucleotides. Base-pairing in nucleic acids shows characteristic signals in the imino proton region (around 10-15 ppm), which are not visible in unpaired nucleobases due to chemical exchange.

In a DNA strand, there are additional energetic advantages of the hydrophobic bases being internalized to the core the DNA double strand and the polar phosphate backbone being more uniformly exposed to solvent.
Thermodynamically, the formation of the two individual molecules of base monomers forming a pair the solvent will still be exposed to the hydrophobic bases. So such a single base pair would be driven by the energy of the two or three hydrogen bonds in the base pair formed.
This is why you will not see such a complex forming in water; the hydrogen bonds with water are at least as strong as the hydrogen bonds with the bases. But such base pairings can in other circumstances be made to form.
If there are no exchangeable protons for hydrogen bonding in the solvent itself, there will be an energetic advantage to the complex.  If there are no exchangeable protons (as with DMSO or carbon tetrachloride) such complexes can be detected by NMR.  Even in cases with single hydrogen bondable protons on the solvent, these bimolecular complexes are observed.


text:
Appropriate statistical test for a student lab?
I am a HS Bio teacher and doing a microevolution lab involving candy. Essentially students use four candy types and push them together until one cracks (Nat Sel). They also do simulations of migration, mutation and genetic drift. They then calculate the "allele frequency" of each type per generation and look for changes. I want to have my students do a simple statistical test to see whether the change was significant or not from each evolutionary factor. I was going to do a chi square test, but someone told me that is not appropriate to this type of experiment.
So, can anyone confirm, is chi square inappropriate, and if it is..
Can anyone point me to a more appropriate test. These are high school kids with no stats background (and I have forgotten virtually all the stats I learned 20 years ago)...

It seems that you're looking for one test that would fit all situations encountered in evolutionary biology. This obviously does not exist. You should provide more information about what you want to test and describe the kind of data you have. And then, I am afraid the question might be off-topic as it is about statistics and not about biology. You may want to try to ask your question on crossvalidated.SE

You may have a point on the off topic bit, though I've already started, so.... :)   To clarify the lab a bit: Students press candies together over three rounds of "natural selection" (starting with 12 candies of each type- m&ms, skittles, etc- 4 types total). Survivors go on to the next round and replicate when the population gets below a certain number. We do another round where kids pour the candy on desks pushed together and some bounce in to the neighbors areas to represent migration, and another simulation for a bottleneck. Students then look at how much each factor affected the #s.

To choose the right statistical method (it is more than just saying "use the t-test") you need to think about your experiment. A good starting point is this figure from Bitesizebio:

There are two relevant articles on that website:

Let’s Talk About Stats: Comparing Two Sets of Data
Let’s Talk About Stats: Comparing Multiple Datasets

Probably also interesting is the definition of statistical terms:

Let’s Talk About Stats: Understanding the Lingo


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How much does the Hayflick limit/telomere length vary across taxa and within humans?
Since they seem to be quite connected, I'm curious if anyone knows of research comparing the Hayflick limit (and presumably by extension telomere length) between different taxa. I've heard the Hayflick limit numbers 40-50 thrown out repeatedly for humans (probably relating to the WI-38 tissue line), but how universal is this really? I Know there are particularly long lived species, but has research been done to correlate this with telomeric length, or do they simply have slower cell division?
I suppose it is worth mentioning that a Hayflick limit of 40 rounds of division is massively different from 50, since each round of division conceivably could double the number of cells...

Many studies, including those of Hayflick himself, have found a strong correlation between a species' Hayflick limit and its maximum lifespan. For example, Galapagos tortoises, which can live over a century, have a Hayflick limit of 90-125, and mice, which live only a few years, have a Hayflick limit of 14-28.
However, the mechanism connecting the maximum number of cell divisions and maximum lifespan is not entirely clear. Cell division is associated much more with growth and development than with the maintenance of mature tissues. Cells that have stopped dividing can be perfectly functional for decades. Cells that don't stop dividing are called "cancer". So, while the correlation is strong, the reason for the correlation remains something of a mystery.
Rose, Michael R. 2005. The Long Tomorrow: How Advances in Evolutionary Biology Can Help us Postpone Aging. Oxford University Press, New York. pp 20-23


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Crossing white eyed Drosophila with sepia eye
So, I'm doing some drosophila experiments with my high schoolers and I had really poor production of wild type individuals for doing the experiment resulting in not enough to cross with everyone... I am trying the following crosses and hoping the kids can still learn from it, but wanted input from any fly-herders on what I can expect with the first two  crosses....
Sepia males x White Eye females
Sepia females x White Eye males
Dumpy males x Wild-type females
Dumpy females x Wild-type males
I don't think the gender separation will make any difference except in the white x sepia crosses since it is X-linked.
What should I get in the sepia x white crosses? Since they both control eye color and are both recessive I'm not sure. Am I correct that w is epistatic, thus should block sepia in an x-linked manner (same as if I had used wild-type instead of sepia)?
I assume the Dumpy x Wild-type will go by the books (F1 all wild-type with F2 3:1 ratio). Fingers crossed.

I posted this on Reddit as well, so credit to km1116 for working this out. Here is his response which follows my original logic (though I'm not sure if he is right):

female x male:
w x se
F1: wild-type females and white males
F2: 1/4 white-eyed males, 3/16 wild-type males, 1/16 sepia males, 1/4
  white-eyed females, 3/16 wild-type females, 1/16 sepia females
se x w
F1: all wild-type offspring
F2: 1/4 white-eyed males, 3/16 wild-type males, 1/16 sepia males; 3/8
  wild-type females, 1/8 sepia females
wt x dp (either direction)
F1: all wild-type
F2: 3/4 wild-type, 1/4 dumpy

My rationale for his answer is the genotype of the cross looks like this:
XwYNN with XWXW nn , where "W" is normal eyes, "w" is white", N is Wildtype and n is sepia.


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What is the name of the appendage that mantis shrimp use to rub their eyes?
In this short clip, you can see the mantis shrimp using some sort of feathery arm/claw to rub its eyes. Looking at a diagram here, it appears possibly that it's using maxillipeds. Someone with knowledge of crustaceans (the behavior is probably not unique to mantis shrimp) would be appreciated.

Mantis shrimp use their first maxillipeds for grooming (maxilliped=modified appendage), which is specialized for this purpose. Details and a picture of the organ can be found in the link. The second maxilliped is their famous specialized organ for striking or spearing prey with enormous force. More about their raptoral appendage, with links to further references. Mantis shrimp also have very complex eyes, described as "...the most complicated visual system of any animal on Earth". Their eyes can see a range from ultraviolet to infra-red and has specialized regions for observing motion, form, depth, and color. More on Mantis shrimp eyes can be found in Marshall & Oberwinkler (1999).


text:
Can anyone identify this bird - small flock in ireland?
I saw small birds in a flock of about 6 or 8.
Here are details that can be seen on my best photo - The beak looks pointed and also the legs are quite long and the eye has a bar across it.

I have a recording - how do I share it??

The body shape and colour reminds me of a greenfinch (Chloris chloris). They also form flocks. However, the beak on your pictures doesn't look very 'finch-like'. Can you tell us anything about the beak shape, or do you have a picture, where it's better visible?

the beak seems to be pointed - the birds were far away - i will try for a closer shot if they return -

the beak looks pointed and also the legs are quite long and the eye has a bar across it-

Maybe edit your question add these additional information there. Not everyone reads all the comments :)

@Marzipanherz You can probably make an answer from your comment.

I also first thought of green finches, but they have a much stronger beak.

@Remi.b Richard's comment about the pointed beak made me doubt. I'm still searching for a better fit.

They're definitely not greenfinch. The horizontal bar across the eye rules that out. I think yellow-crowned wagtail (Motacilla flava ssp. flavissima) is a better fit. See https://commons.wikimedia.org/wiki/File:Motacilla_flava_flavissima,_Kingsbury,_Warwickshire_1.jpg.

@bshane I think Motacilla would have more visible stripes on the wings.

@bli: not always. Worn plumage loses a lot of the distinct scalloping. e.g., https://www.google.com/imgres?imgurl=http%3A%2F%2Fwww.hlasek.com%2Ffoto%2Fmotacilla_flava_da5089.jpg&imgrefurl=http%3A%2F%2Fwww.hlasek.com%2Fmotacilla_flava.html&docid=uMycFdL-fJqLvM&tbnid=7l0k0nvS20RrGM%3A&vet=10ahUKEwjmspmyktzcAhVSQd4KHSL5C3cQMwiHAShEMEQ..i&w=626&h=500&client=ubuntu&bih=1079&biw=1920&q=motacilla%20flava&ved=0ahUKEwjmspmyktzcAhVSQd4KHSL5C3cQMwiHAShEMEQ&iact=mrc&uact=8

Thanks Chris - If they return I will try for a recording - however the Chiffchaff, Willow Warbler and Wood Warbler are described as having black beaks - these have distinct light coloured beaks - perhaps juveniles / male-female differences??

@user45303 - You can make edits to your post using your original account (Richard). You do not need to create another account. Well, if you have forgotten your password then see here what to do. For now, Iam rejecting your edit.

With the new image it is still hard, but I think I identified the bird. I think this is a Willow warbler (Phylloscopus trochilus) which is pretty common in Europe.
The drawing from the Royal Bird Protection Society (source here) is pretty similar.

Unfortunately, there are two pretty similar birds living in Ireland which can be confused with the Willow warbler. These are the Chiffchaff and the Wood warbler. Differences are small, the Chiffchaff has dark legs and a smaller eye stripe (both of which the bird in your image has not), while the wood warbler lives in woods and has a more yellow breast (and is also much more rare). The links provide audio files with the songs of each bird, this should help you making a final decision, as they are pretty distinct.

In the US warblers typical travel in small loose flocks.


text:
Identify a bird by its feather
Gray feather with red tip, about 5 cm (2 in) total length. It was found in Lexington, Kentucky (central western USA).

Where did you find it (which type of ecosystem and what country / region)?

Good point. Sorry, I'm not used to posting here. But I think the answer is below. I'll try to give better information in the future.

It was in our back yard in Lexington, KY


text:
What is the main difference between "Terminalia ivorensis" and "Bucida buceras"?
Google images yields photos of similar trees, but they have separate Wikipedia article. I want to know the major difference allowing to identify which is "Terminalia ivorensis" and "Bucida buceras".

I consider it slightly unlikely that one can find images of Terminalia ivorensis online. The African Plant Database (which started out as West African Plant Database, and has a really good coverage in the area) doesn't have contributed images, and this is a West African species.


text:
Biopython error
>>> from Bio import SeqIO
>>> for seq_record in SeqIO.parse("ls_orchid.fasta", "fasta"):
... print(seq_record.id)
  File "<stdin>", line 2
    print(seq_record.id)
        ^
IndentationError: expected an indented block

First I import the sequence and then whenever i try to print using this
" print(seq_record.id)"
        ^
I get this error
IndentationError: expected an indented block
how can I preent that need help..

This question appears to be off-topic because it is about a bioinformatics software development issue.

basic python question. I suggest a quick search on stackoverflow.com

Biopython is a module of python and the language is structured in the same way.
This particular example does not make it clear how and where to find the files and extract the data you want. First, you need to download the file and put it in a place where python will find it. Below I have an example where I was able to extract the record id. I first set up a fasta file for my code to go through and then it adds the data to a list (seq = []). The "with open" line joins my data files together. The line ends with a ":" to signal to the program that there is a block of something to do in the next line. Indentation is important in python and after a ":" the next line is indented 4 spaces.
from Bio import SeqIO

data_dir = 'data' #folder where my data files are
data_file = 'ampk.fasta' #name of file I want to get data from

seqs = []

with open(os.path.join(data_dir, data_file), "rU") as f:
    for record in SeqIO.parse(f, "fasta") :
        seqs.append(record)
        print(record.id)

Hope this helps!

okay I will try and let you know


text:
SDS Laemmli Gel that did not solidify
In my lab class this week my partner and I were making a SDS-PAGE Laemmli Gel. We made the resolving gel solution which was made of 30% Acrylamide, DI water, 1.5 M Tris-HCL pH8.8, and 20% SDS. We mixed the measured amounts into a conical vial. We then added 10% APS and TEMED and mixed carefully. We then immediately pipetted the solution into the casting stand so the gel could solidify. We then pipetted DI water onto the top of the gel so it would not dehydrate. We left some solution in the conical vial so we knew when the solution would be solidified completely. After waiting 45 minutes the gel in the conical vial was solidified but the gel in the casting stand was still liquid. How come the gel did not solidify in the casting stand but solidified in the conical vial?

How long is your polymerization time? How old are APS and TEMED? Why do you use water on top and not an alcohol?

the polymerization time was said to be ~30-45 minutes. The TEMED was provided to the class so I'm not sure how old it was but the APS was made during the lab. Also the teacher suggested to use DI water so the gel would not dehydrate. Would it be the water that could have caused it to not solidify because that's the only thing that was not added to the conical vial which did solidify

You put a liquid on top of the gel because you want to avoid contact with the air. Air catches the free radicals needed for the reaction. Water can mix with your solution and dilute it (depending on the amount of water used and how cautious you pipetted it), thats why I usually use a few hundred microliters of isopropanol.

The polymerisation time appears to be rather long. From personal experience, it rarely takes more than 15 minutes for both the gel in the falcon tube and casting stand to solidify. How much APS and TEMED did you add?

The water shouldn't be an issue if it's layered carefully.

I would guess that the casting glass may have ethanol from cleaning process, or the water you added diluted mix.
Sorry for not entering this as a comment, for I dont have the necessary rep.

We added 150 ul of 10% APS and 15 ul of TEMED. The water was also added a couple drops at a time alternating between each end of the casting stand.

I don't understand, how do you know that the gel mix did not polymerize in between the glass plates? For example, did you attempt to remove the layer of water so that you could pour the stacking gel and discovered that the entire solution remained liquid? The liquid that you layer on top of the freshly poured gel serves two functions. As mentioned by others it excludes air (oxygen) that would inhibit polymerization. It also helps to form a smooth flat surface to pour the stack. 0.1% SDS works nicely too. Best bet: the plates were not clean enough and inhibited polymerization.

This is very strange. I used the variables given here and yet it solidified much faster. Additionally, I remember the one time I added 50µl of TEMED instead by accident and the gel solidified within 30s after vortexing. I agree with @mdperry's conclusion that the glass plates could have been dirty, or that the TEMED was somehow nonfunctional.

Could be a problem of improper mixing too.


text:
How much do the COVID vaccines decrease the spread of SARS-CoV-2?
The vaccines targeted at COVID-19 are measured against their ability to reduce the occurrence of the disease, COVID-19 (The Vaccine Efficiency or VE).  It does not measure its effect on the underlying virus' ability to spread.  From what I understand, this is the standard approach used for measuring the effectiveness of vaccines.  So called "sterilizing" vaccines, which prevent infection of others, are relatively rare.
From what I have read, it is generally accepted that the COVID-19 vaccines do indeed decrease the rate of transmission of SARS-CoV-2 (if I am understanding properly, this reproductive rate is typically given the label $R_T$, distinguishing it from $R_0$ which is the reproductive rate excluding any proactive behaviors we undertake).  However, I have not found any estimates as to how much it decreases transmission, which is an important number to estimate when projecting the long term behaviors of SARS-CoV-2 in society.
Are there any estimates to the effect the COVID-19 vaccines have on the reproductive rate of SARS-CoV-2 backed by studies?

https://www.nature.com/articles/d41586-021-02054-z cites some Israeli studies stating that vaccinated, infected people are 78% less likely to transmit to members of their household than unvaccinated, infected people ...


text:
How can stem cells be counted in body?
In the study of supercentenarian woman here, it was found that all her blood cells are formed from only two stem cells (ancestor stem cells).
How scientists can distinguish cell ancestors?
Aren't all genes of one body identical?
If there are somatic mutations, which are increasing while lifetime, then how they could know, that there are more ancestor stem cells in childhood? Shouldn't it be less mutations and cell lines less distinguishable?
May be stem cells only can be counted in oldness.
Then how is it possible to know, that there are 20000 of stem cells at birth?


text:
Finding a easy and cheap method for dyeing dNTP
I want to measure OD to know the concentration of dNTP. Any idea for dyeing dNTP at cheapest price and easiest way ?

There may be such a dye, but I'm not aware of it.  The standard way to measure dNTPs, and nucleic acids in general is by absorbance at 260 nm. This article has a table of molar extinction coefficients for dNTPs (Table 10.2). They are
        wavelength      molar extinction coefficient
dATP    259 nm          15,200
dCTP    280 nm          13,100
dGTP    253 nm          13,700
dTTP    267 nm           9,600

In practice for routine work with DNA you can use 260 nm and assume that an absorbance of 1 corresponds to [DNA] = 50 µg ml-1. Using an average of the values in the Table, and disregarding the fact that these are actually at different wavelengths, and using an approximate average dNTP MW of 500, I calculate an absorbance for a 50 µg ml-1 solution of all 4 dNTPs as 1.3, so that's a reasonable agreement given all of the assumptions I've made.
There are fluorescent dyes that can be used to measure DNA, but these rely upon the presence of stacked bases, so are not useful for free dNTPs.
So, bad news if you don't have a UV spectrophotometer. There are plans available for construction of simple spectrophotometers using Lego and a few optoelectronic components. These tend to use LEDs as a light source, but I don't know if UV LEDs are available.

It will be difficult to achieve the level of wavelength resolution required to differentiate dNTPs, considering that you are doing a DIY experiment. Only very precise spectrophotometers can achieve it and they are expensive.
Another technique you can try is thin layer chromatography(TLC). It is cheap and you can easily do a DIY. But you need standards in order to differentiate NTP, NDP and NMP; the latter two might arise because of degradation during the experiment. People generally use a cocktail of anti-oxidants and protectants to minimize the degradation of nucleotides.
A strategy to go about this is to remove the phosphate using phosphatase and then do a TLC with Nucleosides (which are quite stable).
You can check this review on nucleic acid TLC.


text:
Preparation of normal DNA polymerase
i don't need Taq polymerase. Just need normal DNA polymerase. So the main idea is preparation polymerase directly from Lactobacillus without using cloned vector. My idea is electrophoresis using agarose gel and calculating the isoelectric point of Polymerase based on protein sequence to know where they will migrate after electrophoresis. Is it possible for this idea ?

I think it would be easier to use heterologuous expression of a tagged protein, e.g., a His-tag. You could then easily extract that protein from the lysates using a Ni-column. There are many more tags available, but the His-tag is the most common one.

I guess this is still related to your DIY project, you should specify it, otherwise people will assume you are in a standard lab with full access to state of the art equipment. 'Normal' DNA polymerase is not thermostable. It will be inactivated at the first cycle of PCR, that's why most polymerase we currently use comes from organisms that live in very hot environments.

@GianpaoloR I'm running another experiment so taq polymerase for PCR is not necessary now. In this experiment i need to using normal DNA polymerase because normal DNA polymerase can synthesis in room temperature but taq polymerase not. No matter if other method could help me to learning more about biology even it's impossible for me.

This is a very ambitious project for a DIY-er. To explain why, I'm going to use the example of DNA polymerase I from E. coli.
DNA polymerase I is the most abundant DNA polymerase in the E. coli cell, but nevertheless is still only present at around 300 copies per cell

Ishihama Y et al. (2008) Protein abundance profiling of the Escherichia coli cytosol. BMC Genomics. 9:102.

This was the first DNA polymerase that was ever purified, in the laboratory of Arthur Kornberg

(Enzymatic Synthesis of Deoxyribonucleic Acid : I. PREPARATION OF SUBSTRATES AND PARTIAL PURIFICATION OF AN ENZYME FROM ESCHERICHIA COLI Lehman, MJ et al. (1958) J. Biol. Chem. 33:163-170.).

In this paper a method of purification is described that starts with 60 litres of culture. There is also this statement:

"One kilo of E. coli yields less than 10 mg of the purified enzyme."

I'm guessing that you have no experience of protein purification, so I'll just tell you that this is very discouraging: these days, with overexpression, people expect to get mg quantities of their protein from just a few grams of bacterial cells.
This is a clear illustration of why it is pretty much unthinkable for you to set out to purify an enzyme like this using small-scale methods, and explains why techniques of overexpression and the use of affinity tags have revolutionised protein purification.
And it gets worse: even if you did manage to purify some DNA polymerase I from a bacterial source you would find that it is actually not very good at making DNA. This is because as well as extending primers in a 5'>3' direction, it is is very good at degrading DNA in the same direction. In other words as the enzyme moves along a template molecule, copying it, it will degrade any existing DNA that it meets "ahead" of its direction of travel. This is why the most commonly-used form of this enzyme in research is the "Klenow fragment", a fragment of the polymerase that lacks the 5'>3' exonuclease activity. This fragment was originally made by treating DNA polymerase I with the protease subtilisin, but it is now expressed from an engineered polA gene. Klenow fragment polymerase was used in the original PCR experiments (it had to be added afresh at every cycle).
I don't know how much money you have to spend on your projects, but in fact these enzymes can now be bought relatively cheaply.  I don't think that you can expect to make them for yourself, without this becoming the actual project.
Finally, if you can get hold of a strain of E. coli with an expression plasmid for Taq polymerase then you really could make your own very easily - I've done this myself (but I no longer have the strain) and the heat stability of the enzyme makes the purification trivial. It would be much easier to use your own Taq with a heated water bath than to try to make DNA polymerase I. But obviously it depends upon what you are trying to do: Taq polymerase may be unsuitable for other reasons.

Incidentally - I perhaps haven't made it clear that although you don't need mg of enzyme for typical molecular biological applications, it is impractical to purify very small amounts of a protein.

Thank Alan Boyd so much, you always give me a quality answer for my question and give me a direction what i need to do next. I'm planning to create a DNA sequencer, the main idea for my project is heat the tube to 95 oC -> 60 oC for primer can bind. And then cool down to 37 oC for normal polymerase can work. I need temperature room because i think one of my sensor chip can not work in high temperature. I never work with plasmid before and it's seem that i need to get used to it.

I'm an IT engineer and all of my worked money is being spent for research biology. A lot of money was spent and i want to save money as possible for my future research. Enzymes are very expensive for me as the price i see in this table at http://www.enzymatics.com/products.htm . However it cannot make me downhearted.

@ĐứcUltraSoft: if you want to do some research then the best option would be to work as an intern in some lab where you may also implement your ideas. Biological research especially molecular biology is a costly affair and you may end up losing more money.
I think that you can write to George Whitesides who is in Harvard. His lab is into bio-entrepreneurship.

@AlanBoyd do you how many molecule of taq polymerase is needed for a PCR reaction ?


text:
Why not use SDS-PAGE as a method to detect viruses?
Recently, I have been researching about DNA and I know the most popular method for detecting viruses is based on DNA. After learning about proteins, I wonder why we do not detect viruses based on virus proteins?
If it becomes true we can save a lot of money, instead of doing a PCR reaction, we can detect more than one disease in a single reaction.
For example, when I have the sequence of a virus from NCBI Genebank, I can know how many proteins the target organism produces and I can calculate the molecular weight of each protein. Then running it in a SDS-PAGE gel to see what the bands look like, may be that I can know the target organism is present or not?
For example, HIV produces 9 proteins and the molecular weight of each protein is different.
So after doing electrophoresis, I have 9 bands. Based on the distance between each band and based on the molecular weight, it's possible to know HIV is present or not?

Put simply, the answer is that you could seek to detect viral proteins, but because these proteins would be very minor components of your sample, you would have to use an immunoblotting technique to detect a specific viral protein. While immunoblotting is quite a sensitive technique, I think its fair to say that it can be technically demanding. In contrast a PCR reaction is exquisitely sensitive and is fairly robust. PCR is also technically more staightfoward because DNA is generally easier to work with than are proteins.
Before the advent of PCR, immunoblotting would have been the method of choice.

also many viruses will have nearly the same molecular weight and therefore a nearly identical SDS PAGE pattern.

@AlanBoyd point, comment removed.

Absolutely correct.  And to the advantage of "more than one disease" this can, and routinely is done, with qPCR, with a much higher sensitivity that can reasonably be obtained via western or even ELISA (which is a viable detection system often based on protein).


text:
The role of antibodies interacting with cancer
I'm learning about antibodies. As I understand it, antibodies detect stranger cells/bacterial/viruses by the molecules present in their membranes. In cancer cells, the cancer cell have produce some unknown protein, so why antibody cannot detect that molecule and kill them?
Is it that the cancer cell is surrounded by a normal cell?
Or is the protein it produces is not obvious in the cell membrane?
Or that the protein can bypass antibody? if so, why?

This article on the University of Kentucky website is pretty good, it seems the cancer cells hide because they do not express CD80, a co-stimulating molecule. Co-stimulating molecules are often necessary for an effective immune response.
This is a big part of the article and I think they do a good job of explaining it so I'll just quote:

"People once thought that cancer occurred because of a weakness in the
  body's immune system," says Yannelli, "but we have known for years
  that overall that is not the case. The immune system in cancer
  patients actually works quite well. Otherwise, they could not survive
  the disease for long periods of time nor could they fight off other
  viruses, which they routinely do. The problem is not that the immune
  system doesn't work; it is that the immune system doesn't recognize
  the tumor."
According to Yannelli, for the body's defense system to work properly,
  key antigens must be presented in a specific way to lymphocytes, the
  cellular mediators of the immune system, which then kill the cancerous
  cells. Normally, antigens are tipped off by co-stimulating molecules.
  When the co-stimulating molecules are not present, the immune system
  does not "see" the cancer cell, and therefore does not act to destroy
  it.
Scientists have discovered that lymphocytes have very narrow job
  descriptions, each one designed to attack only a specific molecule.
  The researchers have also discovered a wide range of co-stimulating
  molecules. A critical co-simulating molecule is CD80. Because cancer
  cells do not express CD80, they are invisible to the lymphocytes
  responsible for destroying them.

This paper has more on the function and mechanism of CD80.
Thanks to @Nico for pointing out that not all cancers are the same so the generalization discussed in this answer may not be true of all cancers.

We also need to specify that "cancer" is not ONE illness but a series of very different pathologies with very different etiology and development. What is true for one cancer may not be true for another one, so it is always difficult to generalise.

In general the immune system sees the world in very simplistic terms: Self vs Non-Self. Ideally, threats to the health of the animal are all non-self and can be attacked strongly. Whereas self cells and proteins are tolerated - effectively invisible to the immune system. Cancer is a particularly insidious disease because it arises, not from the outside world, but from subtle changes within the host's own cells. Therefore, cancer cells are starting as 100% self, but can adopt changes that make them more different over time.
A good review of this interaction between cancer cells and the immune system can be found here.
A second aspect of your question that is important is how antibodies are made and what they are capable of 'seeing'. I wrote a short essay about B cells, the cells that produce antibodies, and how they are activated on my blog. That might be a good resource to check for you - I also link to one or two other articles there. Briefly, B cells can only 'see' cellular antigens that are either expressed on the cell surface or secreted. Although some cancer cells alter these types of molecules, many more are part of the internal workings of the cell and therefore invisible to antibodies. For this reason, many oncologists focus on T cell responses, which are capable of detecting non-native internal antigens.


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Cancer: what does it mean "at presentation"?
Unclear to me what this means:

"Objective The biological heterogeneity of hepatocellular carcinoma (HCC) makes prognosis difficult. We translate the results of a genome-wide high-throughput analysis into a tool that accurately predicts at presentation tumour growth and survival of patients with HCC."

Reference

https://en.wikipedia.org/wiki/Presentation_(medical)

"At presentation" means "when a symptom/diagnosis is observed/noted by a medical professional".
In the paper you cite, they are doing microarrays on biopsied tumor tissue. "At presentation" here effectively means "at biopsy", at which time that a patient is diagnosed with hepatocellular carcinoma. It's important to note that their sampling is based on presentation time because tumor age at presentation is not going to be the same for everyone - some patients may "present" with symptoms and be diagnosed when a tumor is relatively new, others may present when a tumor is advanced. Presentation is both a convenient and clinically relevant time point - it's the earliest time a prognosis can be offered.

"Presentation" is the event where the patient first visits the doctor.
For example, "the patient presented with yellowing skin and complaining of abdominal pain".
The paragraph you have doesn't make this clear, they assume you have already had your genome sequenced prior to visiting the doctor, who looks at the result, uses this tool and predicts what is likely to happen to you and your cancer based on your DNA.

This answer is a bit wrong; this is not based on prior genome sequencing. They're doing microarrays on biopsied tumor tissue and identifying particular genes for prognostic benefit. Presentation is when the patient is biopsied and diagnosed.

Thank for polite comment, Your answer covers it so I will delete mine tomorrow.


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What are fish seeds?
I stumbled upon this term in a section called Induced breeding of Carps in an Applied Zoogy book.

Traditional practice is to collect seed from the natural riverine habitatwhich also contain uneconomic,unwanted and predator fish species.

What are seeds? Does it mean zygote ?

I can't tell from the quoted text, but could this refer to milt/sperm? 'Seed' (not seeds) is used in this context.

Hmm. This FAO document seems to imply that it is a collective term for all the young life stages of fish from hatchling to fingerling.

Interesting - looks like this is used fairly widely in fish farming but not actually defined anywhere I can see. From the context, another possible definition is 'juvenile life stages used to start a fish farm, usually but not always collected from the wild'.

Fertilized fish eggs are known as Fish seeds. In simple words, they are the baby fishes used for seeding new Ponds in fisheries.
They can be used in 4 stages:

Hatchling : Larvae stage where it have yolk sac hanging below from where it draws its nutrition for 2–3 days. It can't eat from outside as mouth is not formed yet!

Spawn: Stage where mouth is formed.

Fry: About ~2 cm long, now, it can eat small size zooplanktons!

Fingerling: About ~15 cm long. It is now becomes easier to transport these kinds of fishes to Ponds.

Source (Picture Credits) :
Pic 1. https://www.ucdavis.edu (https://www.ucdavis.edu/sites/default/files/images/article/trout_lg.jpg)
Pic 2. Wikimedia (https://upload.wikimedia.org/wikipedia/commons/f/fe/Anemone_Fish_protecting_it's_spawn.jpg)
Pic 3. Fishbio (http://fishbio.com/wp-content/uploads/2011/03/Oncorhynchus-mykiss-fry.jpg)
Pic 4. Deveron (http://www.deveron.org/wb/media/images/parr.jpg)

''Fertilized fish eggs are known as Fish seeds. In simple words, they are the baby fishes used for seeding new Ponds in fisheries.'' -can you support your claim with citations?

A fish 'seed' is a juvenile fish to be used for fish farming.
'Seed' appears to be an industrial/technical term in fish farming used to refer either collectively to the juvenile life stages of fish (hatchling, spawn, fry and fingerling), or specifically to fry (the stage after the yolk sac disappears and before scales form).
It is used in contrast to broodstock or broodfish - essentially, it is what you collect from broodfish, or from the wild, to start a new farming 'batch' (fish farming is a batch process). It sounds like it can be used as a mass noun.
It is surprisingly difficult to find anywhere this definition is stated straightforwardly, although it is used in multiple places in the fish farming literature. However, the definition can be inferred from the context. For example, it is used as a collective heading for the columns "Feeding fry", "Dhani" (a Bengali term for post-larvae) and "Fingerling" in Table 3 (Section 2.1.2, "Broodfish requirement") of the FAO manual on seed production of carps. However, in the FAO document "Seed production: an overview" and this presentation on Slideshare it appears to be used interchangeably with 'fry'. It is clearly differentiated from eggs; for example in the Slideshare presentation above it is mentioned that seed is produced by hatcheries (so consists of hatched fish).


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Need some good resources to learn about Protein function and structure
I'm taking a course on biochemistry at edx.
Since I'm a computer science student, I'm having some trouble in understanding many biochemical concepts. While the first module was just fine, I found the second module really hard to understand. I have problems to learn and understand about amino acids and proteins, so I would like to study apart from some good resources.
Which is a good resource to learn about amino acids and macromolecules?
Also, please let me know if this book is up to date?

Lehninger Principles of Biochemistry is a good text book.

Hello George. It would be much easier for others to answer your doubts if you ask about a specific doubt. There are a lot of things to know about proteins and we cannot know what the second module has that you do not understand. See the tag-wiki on book recommendation. You should address all those points otherwise we cannot suggest a good book as per your requirement. Alternatively, you can ask clarification for your doubts here but make sure that you do not ask too many questions in a single post.

If amino acids and macro molecules are what is troubling you, then just read any biochemistry textbook printed within the last few years.

I personally prefer Berg Stryer Biochemistry alot compared to Voet Voet Prat. There's also the lectures from Kevin Ahern. and there's a book by Ahern as well. As you see these preferences depends on each individual even though a particular text book can be recommended by a course. Another reference is "The Cell" since biochemistry is a study at molecular level and provides indepth explanations on DNA and Protein synthesis.

https://en.wikipedia.org/wiki/Ramachandran_plot

Given your background (not a biologist or chemist) you would probably find the introductory material in a biochemistry "lite" textbook more accessible/useful than a hardcore text for specialists.  As a co-author of a biochemistry textbook, I can tell you that there are essentially three different classes, or types of texts.

Comprehensive textbooks, of which I think Voet & Voet is an outstanding example, are designed for biochemistry majors who will be taking a full year, in-depth course, typically in the 3rd year of their undergraduate studies.
The general introductory texts are usually thinner, and are designed to whip through the same material in half a year--aimed at life-science students in other majors. Some of these books that I described as "lite" are further targeted to, for example, nursing, medical, or dental school students.
The third category of text tries to be multi-purpose, and appeal to both markets. The current versions of Stryer and Lehninger fall into this class (they are both excellent in my opinion); the instructors can cherrypick the readings if they want to use the book for a half-year course.

Voet, Voet, and Pratt (mentioned in the comments above) is a very good diluted version of Voet & Voet. As you can also tell from the comments, different texts have different flavors, and so which one is best depends on the reader.
It sounds like a more general "concept" based book might be useful for you (to provide background and context). The very thick "Molecular Biology of the Cell" by Alberts et al., has a lighter cousin named Essential Cell Biology which would be well worth looking into.
Keep in mind that when tackling a university-level biochemistry course the instructor has to assume that you have passed university level introductory courses in general chemistry and in organic chemistry. Biochemistry majors typically complete full year courses in both these subjects, as well as a watered-down version of physical chemistry.

I want to add a book to the list that may be easier for you since it takes it from the perspective of Physics:
Finkelstein & Ptitsyn, Protein Physics, Academic Press (2002). ISBN 0-12-256781-1.
It covers structure, thermodynamical processes in proteins, mechanism of folding, function, a bit of bioinformatics. It is designed as a master level course in Biophysics, but it takes most things from the beginning: it even explains what the partition function is (but it helps if you know it already). No prior knowledge in biochemistry is required.


text:
Would the human body react faster to touch or sight
I am currently working on an assignment for my physics class, but I believe my question is biology related.
For a two person lab, I am trying to time how long it takes a tennis ball to fall two meters, Using basic physics equations I was able to determine that the ball should take around 0.66 seconds to hit the ground but my results are generally very off.
Would I be able to react faster if I were to stop the timer when I felt the ball hit my other hand, or would I have a worse reaction time than if I were to use my current method of stopping the timer when I see/hear the ball hit the ground.

Try them both and see what works better. Why bother asking here?

Averaging them both would work, but technically speaking I want to know what would work. I am at home and I have no access to the materials .

Pretty certain it is touch. As far as I know the sensations of haptic feedback can create a motor reaction without requiring any higher-order cognitive processing, meaning that they'll take the monosynaptic route. On the other hand, the visual processing takes longer, because in this case, higher order cognitive processing is required.

Are far as experimental design goes, you'd have to take into account that the person stopping the timer is probably predicting when it will hit the ground. They'd be looking at the movement of the ball in the few centimetres before it  actually hits the two-metre mark. Similarly, you'd have to determine how to start the timer. Do they start after they notice the ball being dropped, or do you do a similarly-predictive countdown (e.g. "1, 2, 3, drop").

I can't say what I would do, but I would try very hard to find a non-biological "trick" to measure the time lapse. This may be the purpose of the exercise (or it may just be to try to physically see that the equation works). Just to give you some idea of what is humanly possible, scientists have a "trick" (used at the LIGO observatories) that allows measurement of a change in length (of a 4 mile laser beam) of less than the diameter of the nucleus of an atom). By the way, I agree .. TOUCH.

A "touch" or "haptic" sensation will be much faster due to several reasons:

Haptic feedback can be processed without the presence of any higher-order cognitive processing, therefore meaning that the signals are being processed via a monosynaptic route.
There are short reflex arcs between the spinal cord and the limb (meaning that you can react before you are consciously aware of it)

A visual sensation will be much slower:

The processing of the signal will require higher-order cognitive processing, and this takes time. I read that it takes between 50ms-200ms. (polysynaptic route)

Interesting article about this topic

but does it practically make any difference for an experiment like this?


text:
Forgot to vortex antibody before staining
Ugh. Did an immunofluorescence experiment last weekend, forgot to vortex both my primary and my secondary antibody solutions.
And my final result looks dimmer than it should be. Is it possible that not vortexing contributed to this?
I guess the question is, is vortexing really necessary? It would come down to the fluid dynamics. How long does it take for a drop of antibody to reach equilibrium in a 1000µL Eppendorf tube? Is it really more than just a few seconds? Is vortexing really necessary?

It depends on how much time elapsed between adding the concentrated antibodies to the diluting solution and adding the diluted antibodies to the slide. If it was just a couple of seconds, then I can abolutely see there being a difference. Even if it was longer - say 30-90 seconds, even - there potentially could be antibody gradients in the solution without vortexing. There are a few reasons why. First, many unconjugated primary antibodies that are not intended just for immunostaining are provided in a glycerol solution, with concentrations ranging from 10% or so up to 50% or more, sometimes in addition to a carrier protein like BSA (I used to work for a well-known antibody company, and theirs are in 50% glycerol with 10 µg - 2 mg BSA per ml, depending on the antibody type). The glycerol prevents the solution from freezing solid at -20°C, eliminating ice crystal formation, protecting the antibody protein from degradation and allowing for longer storage times without loss of activity. As a result, however, the solution takes longer to solubilize in PBS or whatever diluent you're using on its own, with no additional mechanical activity.
On the other hand, even if the primary (or secondary) antibody is simply formulated in PBS, the equal diffusion of the proteins to all parts of the tube can take some time. As an experiment, put 1 ml of PBS in an Eppendorf tube, then add perhaps 10-50 µl of a dark dye, such as methylene blue, bromphenol blue, or Trypan blue. Watch it dissociate into the solution over time against a solid white background like a piece of paper. While none of these have the exact same solubility profile as antibodies, it's good enough to show you what happens over the first 30 seconds or so until you can't distinguish the dye anymore.
So, always vortex, even if it's at a relatively slow speed, for a couple of seconds to ensure everything is evenly distributed.

I assume that a small molecule dye will probably diffuse faster than a large antibody. So if a dye takes time to fill the entire volume, an antibody likely needs more time.

@user137 that's exactly my point.

i was taught never to vortex proteins that i wanted to do something useful, but the even mixing (by pipetting or inversion or extended nutation or whatever) of a solution is pretty much required for us to say anything meaningful about any biochemical reaction. In conclusion, the @MattDMO answer is right allowing for local variation in methodological customs.

@MaximilianPress you were taught incorrectly. The only protein I don't vortex is DNase I, used for digesting DNA on an RNA prep spin column - and only because the directions say to not vortex it. Proteins are in general very structurally stable, and a weak force such as vortexing won't damage them one bit, I promise. I have been working with proteins for nearly 20 years, and I've never met a biochemist who said that proteins in general, or antibodies specifically, shouldn't be vortexed. Now, if you're doing native conformation co-IPs, for example, you can of course be a little more (...)

(...) delicate than when working with an antibody for Western blotting or immunofluorescence, but even there the danger is more heat than physical force. Antibodies are incredibly tough and resilient, can often work in a wide range of chemical environments, and in general only need to be treated specially when they're conjugated to light-sensitive fluorochromes. Yes, you don't want bubbles to necessarily form in your solution, but again that's only when you're working in a conformationally-sensitive assay.

Thanks for your answer. I imagine that they don't contain glycerol, since they're kept at -4 degrees. Still, a good thing to consider. I will definitely try the dye experiment.
On that note, how long does it take, if at all, for antibodies to "go bad" at -4 degrees? Could that have contributed?

@amd1972 just a friendly reminder - if this answer addressed your problem, please consider accepting it by clicking on the check mark/tick to the left of the answer, turning it green. This marks the question as resolved to your satisfaction, and awards reputation both to you and the person who answered. If you have >= 15 reputation points, you may also upvote the answer if you wish. There is no obligation to do either.

@amd1972 in response to your question - it depends. If there is sodium azide in the antibody, it shouldn't get contaminated, especially if you're using filtered pipette tips (I never use regular tips anymore). You could check with the vendor to see if they've done any stability studies, or if the lot you have has an expiration date. As long as you're careful with it, it should last at least a couple of years - some can last much longer, others not so much. If you wish, you can add 50% glycerol to it and store it at -20, remembering to label it and alter the dilution factor.

Awesome, thanks. One last question. I made another thread about this: what about the ice cooling step after antigen retrieval that's in most immunofluorescence protocols? I forgot that step too. I've been in school and it's been a long time since I've done these experiments, so, as you can tell, I'm rusty.

@amd1972 not sure about that one - I usually do staining on cells, not tissues, and I'm a little rusty on my IHC.


text:
Finding diploid neanderthal alignment data, looking for ambiguity codes
I'm using Neanderthal alignment data from here: http://www.eva.mpg.de/neandertal/draft-neandertal-genome/data.html
Specifically, the .bam files. I would like to find alignments for neanderthal sequences to human genomes that state ambiguities (such as M or R). However, the data from that site does not have the ambiguities. Is there a way to find such information? Or are there no ambiguity codes in neanderthal alignments?
My goal is to compare diploid neanderthal sequences to diploid human sequences. How do researchers typically deal with this given the limited data they have for neanderthals?

Why would you use the draft genome instead of the high coverage Altai genome?

Is there more than 1 whole genome sequence (besides Altai) for neanderthals? I'm looking for at least 5.

Oh, ok. There is the Denisova genome (a close relative of Neandertals), but other than that, you really have to stick to low coverage (it was not clear from the question how many genomes you need). But if you use low coverage, you should avoid using Altai or Denisova genomes for the sake of comparability.

Ah ok, thanks. I'm looking for at least 5 or more Neanderthal genomes. I guess I'd have to stick to low coverage then if I want that many.


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In induced pluripotency experiments, what happens to the cells that do not become pluripotent?
I am new to this topic, so forgive me if my question is naïve. In an induced pluripotency experiment a few transcription factors are overexpressed in somatic cells and as a result a small portion of the cells become pluripotent (fully or partially), but the vast majority of the cells do not. My question is this: what does happen to these cells (the ones that do not become pluripotent) in terms of gene expression levels? Do gene expression profiles of these cells remain unchanged (the same before and after overexpression of the pluripotency factors)? Or do they change and become something that is different from both pluripotent cells and the original somatic cells?


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About sandwich ELISA
In sandwich ELISA the Fc region of primary antibody bind to the polystyrene coated well. But what are the specific interactions (like 'hydrophobic interaction' or 'van der waals force') happen between well and Fc region?

This page from Cole-Parmer includes everything you could possibly want to know about the adsorption of biomolecules to polystyrene. To summarize:

binding is neither covalent nor ionic, but
it is generally mediated by non-specific intermolecular van der Waals interactions, which generally are about 100X weaker than ionic or covalent bonds
the exception is some hydrogen bonds, which can be 10X stronger than the other types

Intermolecular attraction forces are based on intramolecular electric polarities of which two types can be distinguished: alternating polarities (AP) and stationary polarities (SP), i.e. dipoles.

these interactions can be both hydrophobic and hydrophilic, depending on the surface chemistry of the polystyrene

If you're interested in the specific chemistry of everything, feel free to read the entire thing, it's quite detailed.
I would to note one thing from your question. In the absence of a specific binding partner like Protein A, antibodies bind to polystyrene completely non-specifically, meaning that not only does the Fc region bind, but Fab as well, including the CDRs (antigen binding domains). An ELISA depends on the fact that not all antibody molecules bind this way, so many or a majority of them have their antigen binding sites available to capture their target.


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What is the difference between intials cells and meristems cells?
Generally plant tissues:

meristems tissue: meristems cells, apilal meristems, lateral meristems
Non meristem tissue: dermal tissue, vascular tissue, ground tissue

my question:
What is the difference between intials cells  and meristems cells? Are intials cells the same as meristems cells?


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Can a non-response to a vaccine be tested?
The BioNTech/Pfizer vaccine against SARS-CoV-2 is said to be 90% effective.
Is there a test to establish the (degree of) success in individuals of a vaccine at provoking the desired immune response? Or will we all have to hope for the best?

Some measure like this are listed as secondary or exploratory endpoints in the Pfizer protocol. It's a bit technical, but with some careful reading and googling, you should find what you're looking for in the protocol here

I suppose the follow up question is: is per-person efficacy verification likely to be standard protocol in a vaccination program of such a scale?

You can test for antibody titers (and also connected memory) for the persons who got the vaccine and then decide, if these antibody levels are protective or not. Problem with this method is, that it is not clear, what levels are protective. There is still no experience with it for SARS-CoV-2, this is routinely done for other diseases.
According to clinicaltrials.gov, the study design is a placebo controlled trial triple blinded (Participant, Care Provider, Investigator) which enables the direct comparision of the vaccine arm with the placebo arm. Here you analyze the difference between these arms. If the vaccine delivers a sufficient protection, less (or ideally no) people in the vaccine arm get sick - which is exactly what was observed.
A last possibility would be to challenge people, who received the test vaccine with the virus and see, if they develop the disease. However, this approach is highly unethical since you possibly infect a large number of people with a potentially deadly virus, when the vaccine does not work. I doubt that a researcher who wants to do this will get clearance for these experiments by the internal review boards involved in these trials.
This is done in earlier stages in animal experiments, to challenge the immune response.


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Isolating colonies in a pure culture
I want to conduct antimicrobial susceptibility tests for which I've to culture bacteria first. Since I've bought KWIK-STIKs, (KWIK-STIK contains a single lyophilized microorganism which have to be rehydrated before culturing them on agar plates) I referred to this video for instructions.
In the video, the bacteria are rehydrated and streaked on the agar plates to obtain individual colonies. I know one should isolate colonies in order to obtain a pure culture. But why should you isolate colonies if the culture was already pure?
Here, I'm assuming that companies sell bacteria which are already isolated and sampled, therefore are pure (that's why I wonder why isolating colonies is required), but I'm not a professional in the microbiology field (I'm a high school student) so any basic clarification would be welcome.

Your question is not stupid, but it is not clearly expressed. Please rewrite it. Start by explaining clearly what you are doing. Colonies of what? What kind of culture. What is your aim? You refer to a video, but nobody here knows anything about it, and giving us a link is no good because questions here have to be self contained. So you need to carefully report what the video advises, quoting the advice verbatim. That’s how science  works.

@David Ok, I've rewritten it...is it clear now?

Yes that's better. Someone with the appropriate background will have something to go on now.

The answer is that it allows you to check viability of the stock primarily, and thereby also have a population that you know is growing. Bulk cultures from liquid will always contain a proportion of non-viable organisms as there are no easy ways to separate these from the viable.
It also allows you to check to see that the preparation hasn't been accidentally contaminated (or tubes swapped etc.) by another morphologically different colony former - having the right type of organism is very important experimentally. If you wished to check the morphology of the organism by either morphological (i.e. microscope + stains) or biochemical tests, both of these need to start with a pure culture. It is hard to get enough bacteria for most of these tests unless you pick them from colonies - liquid culture just isn't concentrated enough, and can be heterogenous if a mixed inoculum is (accidentally or otherwise) added.
Another reason is that there is some heterogeneity in the culture anyway, no matter what you do to characterize it after growth. Isolating from single colonies, which should be derived from single colony forming units, allows you some control over this population for your bulk preparation, whether this be growth under certain conditions (e.g. antibiotic resistance) or growth rate.
It for instance may allow you to select a large colony, which might indicate a slightly higher growth rate for that founder. If you have something that is grown on selective media, then it may be that a portion of a bulk culture has lost the characteristic for growth on this medium, which means that when you inoculate from the bulk, what you put in might not be what you expected. This can lead to deviations in expected generation time calculations and similar physiological or phenotypic measures.


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Can anyone identify these mushrooms?
Found in the Dolomites, Italy

in a wooden fence:

The first one is dyer's polypore or rusty gilled polypore.
https://en.wikipedia.org/wiki/Gloeophyllum_sepiarium
The second one looks a bit like orange peel fungus, although it could be something different.
https://www.wildfooduk.com/mushroom-guide/orange-peel-fungus/

Thank you! The second one is really similar to an orange peel fungus, unfortunately I did not take more photo that might have helped with identification...