Science topic

Ribosomes - Science topic

Multicomponent ribonucleoprotein structures found in the CYTOPLASM of all cells, and in MITOCHONDRIA, and PLASTIDS. They function in PROTEIN BIOSYNTHESIS via GENETIC TRANSLATION.
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Dear Colleagues:
I have multigene sequences of my target species(18S and 28S ribosomal genes) for a phylogenetic analysis with GenBank sequences of other taxa to see the relationship among related families. The result using each gene looks ok, and now I am trying to do additional analysis based on the sequence concatenation. The problem is that the GenBank sequences of different genes were generated separately using different individuals by different researchers, even though they are from same species. My question is, therefore, is it ok to make concatenate alignment of the sequences from different individuals for my research purpose?
Thank you.
Sincerely yours
Jeongho Kim.
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I guess NO!
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Hello all,
I have been trying to grow E. coli cultures for the past few weeks but every time I extract total RNA, it does not show the 23S and 16S rRNA bands on the agarose gel. I have tried multiple times and reduced the incubation time from overnight to 6 hours. I can't tell if my E. coli is in lag phase or if I have contamination causing RNA degradation. Any help is appreciated.
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Yes, you obviously have RNA degradation. Almost all your RNA is tiny, below the ladder. This should not be so. You have a massive source of RNAse somewhere. Look at your process. It's big. It should not be hard to find.
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In someone who gets a bivalent mRNA shot, what happens, exactly, when the mRNA for BOTH subvariants of the spike is active in the same cell? Such cells are NOT going to just produce two kinds of spikes (Wuhan and Omicron).  They’re going to produce hybrid spikes using various combinations of the proteins that normally combine to form the spikes of the two subvariants.  The spikes are each made of many subunit peptides and proteins that are generated by ribosomes from the mRNA and THEN self-assemble to form the subunits containing three ribosome-generated copies of each spike subunit protein that then have to assemble together to form each spike.  So does this mean that such cell will “normally” turn out a large number of DIFFERENT spike proteins? It does.
“The total length of SARS-CoV-2 S is 1273 aa and consists of a signal peptide (amino acids 1–13) located at the N-terminus, the S1 subunit (14–685 residues), and the S2 subunit (686–1273 residues); the last two regions are responsible for receptor binding and membrane fusion, respectively. In the S1 subunit, there is an N-terminal domain (14–305 residues) and a receptor-binding domain (RBD, 319–541 residues); the fusion peptide (FP) (788–806 residues), heptapeptide repeat sequence 1 (HR1) (912–984 residues), HR2 (1163–1213 residues), TM domain (1213–1237 residues), and cytoplasm domain (1237–1273 residues) comprise the S2 subunit (Fig. 2a) [13].”
We can confirm in Fig. 2a of this peer-reviewed article clear confirmation that the spike is NOT created in one go by a ribosome reading and connecting the 1274 aa (amino acids) in sequence.  Rather, these proteins and peptides are made from S1 and S2 proteins, which in turn are made of NTD, RBD, FP, HR1, NR2, TM, and CT proteins.
There are mutations within (that is, differences between) the genetic sequences of at least each of these proteins: NTD, RBD, FP, and HR1 between tozinameran and riltozinameran or famtozinameran.
So with 2 options for each of 3x4=12 locations, we have 2^12 - over one thousand combinations.  We don’t know how each of these thousand different spike proteins will act in the human body.  We have no idea.  It’s likely they would all be created, but we don’t know.  (I speculate that maybe some combinations wouldn’t self assemble, or would assemble into something totally unexpected.)  Yet we’re (indirectly) injecting them into billions of people.  They have only existed since this bivalent vaccine started being used, and ~99.8% of them have not been studied at all.
What am I missing?
[edit: Edgar, your “answer” below demonstrates no understanding of the question and makes no attempt to answer it – the actual question I asked. I’m asking a scientific question. Please don’t use it as a soapbox for proselytizing. I'm a fan of the mRNA therapy platform - and this wrinkle is concerning. I’m looking for an answer that shows expertise in protein assembly.
  • Also, technically, you’re spreading misinformation as, for example, it is inaccurate to claim that the bivalent vaccines have been through reported clinical trials. They have not. (If that changes in the future, do point to it at ClinicalTrials dot gov or point to the literature if there’s one I’m not aware of.) Also, “no evidence” you say? Heck, *according to Pfizer*: “Myocarditis (inflammation of the heart muscle) and pericarditis (inflammation of the lining outside the heart) have occurred in some people who have received … Pfizer-BioNTech COVID-19 Vaccine, Bivalent. The observed risk is higher among adolescent males and adult males under 40 years of age than among females and older males, and the observed risk is highest in males 12 through 17 years of age. In most of these people, symptoms began within a few days following receipt of the second dose of vaccine. The chance of having this occur is very low” <sic>
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It is not accurate to say that people who have received a bivalent COVID-19 mRNA vaccine (such as the Pfizer-BioNTech COVID-19 vaccine) are at greater risk of harm due to multiple copies of different spike proteins in their bodies. The spike protein is a vital component of the COVID-19 virus and is what enables the virus to enter human cells. The mRNA vaccines developed to protect against COVID-19 contain small pieces of genetic material (mRNA) that encode for the production of the spike protein.
The mRNA vaccines work by prompting the body to produce copies of the spike protein, which triggers an immune response. The body recognizes the spike protein as foreign and produces antibodies to attack it. This immune response helps to protect against future COVID-19 infections. The mRNA vaccines do not contain live viruses, so they cannot cause COVID-19 infection.
There is no evidence to suggest that the presence of multiple copies of the spike protein in the body following vaccination with an mRNA vaccine is harmful. The mRNA vaccines are safe and effective in clinical trials and have been authorized for emergency use by regulatory agencies worldwide.
It is important to note that all vaccines, including mRNA vaccines, can cause side effects in some people. These side effects are typically mild and resolve independently within a few days. However, you should seek medical attention if you experience severe side effects after receiving a COVID-19 vaccine.
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Hi everyone,
I am looking for a ribosome complex on an mRNA .PDB file to visualise the interaction in the PyMol. I was so far unsuccessful at finding it in PDB or Uniprot databases, but want to learn more of how mRNA interacts with the ribosome.
If you have this file, I would be grateful if you could please share it with me.
Thanks,
Maria
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Hi, Maria
4V6I is a ribosome with an mRNA, for example.
Regards,
Alexander
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The popular IRES from HCV or EMCV was used in quite a few plasmid. I am wondering any one try them in fish or bird?
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Yes it works, because fish belong eukaryotes. IRES has role in translation process.
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Hi ResearchGate friends, I have found some visible bands between 18s and 28s rRNA while running the formaldehyde-denaturing agarose gel on total RNA (please check the attached images). This is seen in my experiment group, not in the control group, though the total RNAs in the two groups are co-extracted using the same method.
Anyone can kindly give me a hint if they are precursors or fragments of rRNA? Thanks!
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Hi Matt Wu ! Yes, we did and saw the same thing. Please find attached the bioanalyzer results. The RIN is 8.7.
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I was trying to recalculate the RIN value using the Agilent Bioanalyzer. because it couldn't calculate the RIN value due to 'unexpected ribosomal ratio' error flag. I checked the setpoint for the ribosomal ratio for RIN calculation and it was 0.7. I then edited the fragment size of the 28s and got a ratio of 1.1 but still the error says the same and no RIN value. So, does that mean it is impossible to recalculate the RIN value? Thanks in advance for your kind help.
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I have the same issue with the bioanalyzer, where did you change the threshold to recalculate the RIN value, it only lets me modify the lower marker but the RIN stays the same. And besides, it doesn't detect the peak from the 28S, but the graphics do.
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Text book always said that most important job of RNA is protein synthesis - assembling amino acids into proteins. Messenger RNA carries instructions from nucleus to ribosomes. Transfer RNA transfers each amino acid to the ribosome as needed by the code of the mRNA molecule.
However, it seems that it never mentioned the metals in the protein.
May I miss some knowledge in this field?
Where are the metals?
How the RNA to decide the metal center in the protein?
Any hints?
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Thanks for your reply.
I am not sure if my question is clear. What I need to know is the factors that contribute to the selectivity of the metal center in metalloenzymes and the role, if any, the corresponding RNA/DNA plays in this process.
In other words, is the metal center first, then the amino acids -controlled by RNA to fit its structure to form the metalloenzymes, or if the amino acids -controlled by RNA first and then select the metals to form a stabilizer protein?
If it was the second case, the metal availability in the cell is important, which is always linked to the environment. However, the metal center seems quite stable and it does not change for a metalloenzyme.
I prefer the first case. However, I do not have any references to support my idea. Any hints are welcome
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Running a sucrose gradient and I am only interested in the disome peak, would also like to separate out the monosome peak. Not interested in polysomes past this.
We usually run a gradient 10-50% sucrose to get the full range of ribosomes - from subunits to the high polysomes.
Should I attenuate the glucose density gradient for this more narrow focus? Would 10-30% be reasonable?
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Hey Aditya,
when I was running ribosome profiling from mammalian cells, the 80S monosome and disomy peaks popped up approx. in the middle of 10-50% sucrose gradients (SW40 rotor, 35 000 rpm, 1 h 40 min), so 10-30% sounds like a good starting point to me.
Do you have equipment for fractionation and simultaneous OD measurement?
If yes, it would be easiest to give it a test run to be on the safe side :)
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Hi everyone,
I performed ChIP-seq analysis on E. coli cells using antibodies against one of a transcription factor. During data analysis, I validated ChIP samples against corresponding "input" samples (sonicated DNA, without immunoprecipitation). After sequencing, I got a lot of sequences encoding tRNAs and ribosomal subuints. It is doubtful that the analysed protein interacts with sequences coding tRNAs. What could be the problem? I have to add that during sample preparation I used an excess of RNAse for DNA sample purification. An external company performed library preparation and sequencing.
Does someone have a similar problem with ChIP-seq data analysis?
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Did your experiment include an IgG control or similar? This would help with interpretation of this result. Definitely an odd one!
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I am preparing to design primers to clone a Leu2 maker from a plasmid into Saccharomyces cerevisiae (BY4742) in place of a native gene via homologous recombination. However, I struggling to get my head around the genetic workings of the gene orientations of both the Leu2 gene (from plasmid PAG305GPD-ccdB) and Htz1 from (BY4742). From what I can see, both the Leu2 and Htz1 are coded for on their respective negative strands.
As I understand it, transcription always takes place 5'-3 'direction against the antisense strand, which in this case would create mRNA strands with this kind of sequence 5- XXXXXXXXGUA-3'.
My confusion is this - How is it then translated into a protein if the 'AUG' recognition codon is in reverse. Can a ribosome move from the 3'-5 'end of a strand or is it rather that in this case transcription takes place against the sense strand? Or am I misunderstood something fundamental along the way?
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Fundamental misunderstanding, I'm afraid. Though it's great that you're thinking these things rather than just assuming you know what's going on.
DNA is always transcribed in the same direction, it's just that DNA has two strands, which run in opposite directions.
There is no difference between a gene going 5'-3' on one strand, and a gene going 5'-3' on the other strand, even though those two strands are wound together such that each runs in the opposite direction to the other.
A gene can be encoded on either strand of the DNA, as here. The RNA polymerase copies from one strand, and the other strand in that region is ignored. RNA polymerases only 'see' one strand at a time, and that's the strand they bind to
(slightly confusingly, they bind to the non-coding strand, and they travel in a 3'-5' direction, because they are producing a complementary sequence, i.e. a copy of the coding strand, which consequently runs 5'-3').
The resultant mRNA is single stranded, is a copy of the coding strand sequence (with U in place of T, obviously) and runs 5'-3'.
Note that genes rarely overlap between strands (i.e., HTZ doesn't overlap with PDB3) because this would require a region of shared anti-parallel coding sequence (which is both much more vulnerable to mutational change, and a lot more challenging to evolve). Some viruses do in fact have multiple reading frames that overlap on antiparallel strands, because viruses are awesome, but this is because viruses are under huge selective pressure to have small genomes, while higher eukaryotes have essentially zero pressure for this (the human genome is like 90% filler sequence, for example).
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I need to find a Mammalian cell line that can translate mRNA of a bacterial gene
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Any mammalian cell will translate an mRNA provided it has a good ATG (Methionine) translational start site, usually. A/GXCATGG/A, also known as a Kozak Sequence. What is important is a eukaryotic transcriptional promoter to express the mRNA in mammalian cells. Once you make the mRNA, the ribosome does not know whether the coding sequences is bacterial or not. So any mammalian cell can translate a bacterial gene provided you construct the expression plasmid appropriately.
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I would like to do an MD simulation of ribosomes. However, I do not want to simulate the entire ribosome because of limited computational power.
I want to simulate a spherical subsection of the ribosome and restrain all the atoms outside the sphere. Is there any way to do this?
I am using Gromacs for the simulations.
Thanks in advance.
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Aashish Bhatt I actually want to do AA only in this small subshell.
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Dear members,
I would like to ask for your opinions about the process of yeast cell aging.
Which theory is in your opinion more trustful for explaining yeast cell string?
Thanks for an interesting discussion.
Regards,
Marco
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Marco Eigenfeld i always wanted to undersstand about is yeast cell antiaging and came across your question and am sure the link am sharing is quite helpful for you and your research work.
Basically cIame across this research and the yeast replicative aging is thought to be comparable to agigng phenomena observed in quite a few particularly asymmetrically cells which are already divided or dividing.
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My goal is RNA extraction from yeast's ribosome; still now I made this with TRIsure but the result is terrible. Can someone help me?
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Martina Colasante when we talk about DNA extraction of rhizome
I came across this link and felt might be of great help to you and your reearch
This is basically about the the yeast total RNA isolation protocol that actually works very well and is quite reliable, Ti might take some hours or days to start and if concentrated it is a game changer for sure
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Hi All,
I have a protein sequence cloned in pET21+ vector.
Below is the part of the sequence starting from T7 promoter to the ATG (of my protein):
TAATACGACTCACTATAGGGGAATTGTGAGCGGATAACAATTCCCCTCTAGGATCCTCTAGAAGGAGGACAACCATG
Protein is not expressing at al, I am using BL21 DE3 for the expression. I am only curious to know that does this sequence have Ribosome binding site/SD? Or is this sequence correct for the expression of my protein?
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Your RBS is AGGAGG and the distance between the ATG and the RBS is fine. I would suggest you look elsewhere if you are having problems since the sequence seems to be ok.
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I am doing polysome fractionation for stress conditions for mammalian cell. it seems that peaks of 40S and 60S disappear during stress condition compared to control. I am wondering whether all ribosome are struggling with translation since our translation analysis show us there is an increase in translation.
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Perhaps the answer is a little late, but there are many researchers who may need your valuable question and may also find answers to advance their respective investigations. There are related jobs in:
Analysis of translation using polysome profiling (https://academic.oup.com/nar/article/45/3/e15/2972201)
Studying the Translatome with Polysome Profilinghttps://link.springer.com/protocol/10.1007%2F978-1-4939-3067-8_4
Polysome Profiling without Gradient Makers or Fractionation Systems ( 10.3791/62680)
regards
Reinaldo
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  • His-tagged protein immobilized a  on Ni coated NTA chip.
  • Both the reference channel and ligand bound channel shows binding with analyte (ribosome)
  • negative RU in the resultant curve.
  • Reference channel is Ni free.
  • Running buffer contains hepes,NaCl,EDTA,Mg(OAc)2
  • Both the ligand and analyte are prepared in the running buffer.
  • Reference channel was tried to be coated with BSA and still it's showing non-specific binding.
  • Tris buffer as running nuffer gave the same non-specific binding but the resultant curve gave positive RU. As suggested in some paper I am using Hepes Buffer now. If anyone has any idea please suggest me a way.
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Dear Mariana Amaral, it was an experiment I was trying to do a long time ago. Unfortunately, I had to move on to other experiments as it was taking a long time to standardize. However, the main problem I was facing was the magnesium ions in my running buffer as far as I can remember. Magnesium being a divalent cation was probably binding to the NTA chip surface, giving negative RU. I also had cobalt ion in the buffer containing the his-tagged protein. Besides, ribosomes being such a big molecule needed more standardization which I didn't pursue. However, as far as I can remember, all the above suggestions from other people help understand the crucial points one needs to follow in order to standardize the experiment which I left inbetween.
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We know about that ribosome is composed of proteins and rRNA and rib is the precursor of proteins for ribosome composition the proteins is synthesis by which?
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The proteins that are part of a ribosome are also made on other ribosomes. Remember that in cells there are many thousands of ribosomes so there will always be some functional ones.
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During Replication, helicase, primase, DNA polymerase and ligase proteins travel from wherever they are in the nucleus to a specific region of the whole DNA. Why and how do the proteins reach till the 5 prime end?
During transcription, RNA polymerase travels towards the promoter region. How does this movement happen?
During translation, ribosomes travel towards mRNA and tRNA travels towards ribosomes. How?
Is the movement of protein/ chains random? or is it precise?
If it is precise how does the polymease/tRNA know where the DNA/ribosome is and how does it move in that direction?
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Maybe add the concept of turnover of proteins and the concept of time. When proteins are made, it is taken in vesicles to RE and very occasionally, to other sides of the cell. This happens as it is made. There are proteins which guide the machinery with more proteins to the position indicated by a stimulus. This means the proteins stable the region and start to replicate. This stimulus ( and the replication means mitosis and not a temporal protein) signalling ( and normally CA+2) means this region of the DNa opens and starts the replication. These signals come from the membrane (kinases). The nucleus membrane have holes. The replication of the DNA means nucleus disappears. This promoter have enhancers of replication. There are guides to the zone and part of the answer is epigenome.
Another form of guidance is the peptide signal. Not all mRNA is the coding protein. This means some of the protein does not translate. Maybe this could be exons or Exon shuffling. Why do different exons give different genes. It is not a question in the genome,but the combination of genes which turn to coding areas or ORF (Open reading frame). This means we could cover from virus or jumping DNA. tRNA converts the section of DNA to amino acids but animals need these amino acids whilst plants make them. From the ribosomes the proteins are taken to the RE normally. A very specific protein must be for not to be taken to RER ( and then into vesicles). The glycosylation starts and then turns into a specific protein. It may be random but it must be specific to this place. There are diseases of autophagy. The peptide signals takes it to the ribosomes. The proteins can be taken by the cytoskeleton in chains by miosin and actin. (Some of the proteins although we can say most of them). The u uracyl is specific to the ribosomes. They match together at the same time (or almost at the same time). tRNA match the amino acids with the triplet of the genetic code.
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Prokaryotes take aid of Shine–Dalgarno sequence to detect actual translation initiation AUG among the many AUG codons in the m-RNA. Is there any such kind of sequence in eukaryotes that help ribosomal machinery to detect actual initiation AUG among the various AUG codons? Or is it the case that there are no any AUG codon before the actual translation initiation site in eukaryotes so that recognition sequences are not essential for translation initiation? Or there is different kind of initiation site recognition mechanism?
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Hi,
In eukaryotes there is no need for such a sequence because the initiation complex recognizes the 5' end of the mRNA by the presence of the modified Guanosine cap, then the complex scans the mRNA to find the first AUG. In bacteria, the need for a Shine-Dalgarno sequence is explained because the transcript (operon) contains a number of genes, and every gene should start by the Shine-Dalgarno sequence to mark the beginning of translation.
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Hi,
When using the lambda red system, after removing the antibiotic cassette, scar sequences are left behind. Do these sequences vary from clone to clone? Do they contain stop codons? In the three phases?
This is importart when deleting a bacterial gene that is in an operon with translational coupling: it is essential that the deletion does not insert a premature stop codon, because this would cause the ribosome to fall, and the downstream gene would not be translated. In translational coupling, the first gene´s mRNA UGA must be juxtaposed to the second gene´s AUG (so that the translating ribosome can turn around from the TGA to the ATG).
And if anyone knows of simple systems for making seamless/scarless deletions in K. pneumoniae that would be great!
Thanks for your answers.
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Whether or not stop codons are left will depend upon the specific cassette you are using. The original KD3 and KD4 cassettes do have stop codons in all frames whereas I think KD13 does not. Also many other different cassettes have been made to use, so it depends upon the cassette. However I would not be so worried about it. Many genes in an operon are not translationally coupled, if an ORF still has a good RBS then the upstream translation (or lack thereof) may not have much effect. Likewise transcriptional polarity is not going to be an issue unless you have a long untranslated region.
But you can design your own cassettes so that it leaves behind whatever sequence you wish (plus of course the FRT site).
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I shined uv to plants and got a kegg figure for ribosome pathway after RNA-seq. But what do these components circled in red and blue mean? thanks
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Annemarie Honegger Thanks for your prompt response. Actually, I had ribosome KEGG pathway in a JPG.file format, where I was unable to decode the information. But, now I've browsed (https://www.kegg.jp/kegg-bin/show_pathway?lpb03010), and got some details e.g. /enzyme/protein codes and names etc.
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I am using BSA as block. solution and I think it is not so good to block protein up to 20kDa by mixture of BSA with more than 70kDa. Can you recommend me some better, smaller solution please?
Thank you!
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Of course I agree with dear Adam B Shapiro
I would use a commercial set, you can look for it from suppliers on the market.
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I constructed a synthetic DNA library (scFv, VH-VL orientation) with a 3' reporter and 8x histag. I cloned and expressed this gene following which I performed an ELISA. The ELISA results suggested binding activity of some of my scFvs against their target antigens. I sequenced the DNA of these 'binders' and the results were unexpected. I could detect the full and correct reading of the VL sequences but interestingly the upstream VH sequences had 2-3 stop codons interspersed and were not recognized as correct IgVHs by bioinformatics.
Can anyone please explain to me what may be going on here? A stop codon upstream should usually truncate the protein at that point but the ribosome seem to have read through the stop codons.
I guess what I am interested in knowing is whether the VH is aberrant and non-functional and whether the binding activity I observed with ELISA was contributed solely by functional light chain, VL.
Here is one of those sequences (VH, linker and lambda VL).
CCATG GCC GAGGTGCAGCTGTTGGAGTCCTGGCCGCAGTGAGGAAGGGAACTGACTGCCGAGGCCGGAGGGCAGCCGGCCTGCCGAGGGGAACAGCTGGATCTGGGTCCAGGTCATGGGTAGGTGGGCGCGCAGGCAGACTTTCGTGATGTTGTGCACTTGAGGTGTTGTGCTGGACTTCCACAGCAGACAGGAAGGGTGAACAGAGCAGCCCTTCAGAGAGAAGCACTGGGCAGTCACAGGGATTTGAGGGAGGGTGGAGGACACCCTGCAAGCCCTGCGTGGCCACCAACAGGCAGCGCCCTGCAGCAAGGGGGCCACCGTCTCCTCA
GCCTCCACCGGATCCGGTTCTGGTAGTGGTGCTACTTCTGGATCC
TCCTCTGAGCTGACTCAGGACCCTGCTGTGTCTGTGGCCTTGGGACAGACAGTCAGGATCACATGCCAAGGAGACAGCCTCAGAAGCTATTATGCAAGCTGGTACCAGCAGAAGCCAGGACAGGCCCCTGTACTTGTCATCTATGGTAAAAAAAACCGGCCCTCAGGGATCCCAGACCGATTCTCTGGCTCCAGGTCAGGAACTACAGCTTCCTTGACCATCACTGGGGCTCAGGCGGAAGATGAGGCTGACTATTACTGTAACTCCCGGGACAGCAGTGGTAAGAGTTGGGTGTTCGGCGGAGGGACCAAGCTGACCGTCCTA
PS: I sequenced 2x with different primers and in forward and reverse.
Thank you all in advance.
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Randy, if you're planning to express your construct in mammalian cells, the stop codons will surely do their intended job. In addition, you won't get high expression yields (if any) due to codon usage bias. It's highly suggested to have the cDNA synthesized for mammalian use, most gene syntesis companier are offering free online tools for that. It's not that expensive, should be much less than 1000 USD for a ScFv now. This will save you a lot of nightmares and frustration. Don't forget to add the usual culprits (a promoter like CMV, a Kozak consensus element for a strong initiation of translation. Replacing the stops with what they actually have been translated to by the permissive bacterial strain) and replace the pelB bacterial signal sequence with sth that works for Eukaryotes, e.g. the IgG-kappa signal. Have a look at the pSecTag2 vector maps e.g. they will tell you what you need.
If your ScFv turns out to be truncated, maybe replace the stops in question with sth innocent (Gly, Ala, Ser Thr or similar).
Good luck!
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I am running a simple experiment with a co-culture of bacteria (P. aeruginosa and an E. coli strain) in LB to study their interactions with regard to biofilm formation. Is there a good or feasible way to separate these two bacterial species from this co-culture in order to separately analyze their respective ribosomal profiles?
Thanks!!
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I do not see the difficulty in separating Pseudomonas aeruginosa from Escherichia coli culture. From pigmentation alone, the cultures are different, just pick and perform separate quadrant streaking on LB agar and subculture until pure cultures of each is obtained.
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I actually have several questions:
1) Where do I go to learn about the physical chemistry of the interaction between ribosomes and RNA made with pseudouridine or 1-methylpseudourine? 2) Are there any concerns that a DNA vaccine will integrate into host genome in the complete absence of any sort of integrase? 3) Is an immune reaction against ordinary mRNA a problem? Are there any papers documenting instances of this? 4) My biggest question: how is an mRNA with modified nucleosides, a 5' cap, and poly(A) tail manufactured in the quantities necessary to inoculate an entire nation?
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Hi Laura,
Original human vaccine for rabies was unmodified and caused rather severe inflammation due to overstimulation of pattern reognition receptors like tlr3, 7, rigi mda5 etc. So the approach taken to make the vaccine acceptable was to incorporate pseudoriridine to dampen the response.
Rna requires a reverse transcriptase to convert it to dna first. Unless the cell is infected with a retrovirus this is highly unlikely as it violates central dogma.
You use a plasmid and transfect a cell line and harvest mrna. Perhaps they ate using oligo synthesizers to get the correct ratio of pseuduuridine incorporation.
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Hello everybody!
so I have a plasmid which has a synthetic intron (IVS) & an internal ribosome entry site (IRES), and I'm thinking to remove them as I'm cloning due to some matters related to preferable enzymes restriction sites, and my question is :
What will be the effect on the expression of my gene of insert after I remove them?
looking forward to answers and thank you already !
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Assuming your gene is protein coding:
Removing the IVS will probably lower the expression of your protein.
Deleting the IRES could cause major problems, it's there so two different proteins can be translated from a single mRNA. This could result in cells not expressing the selection marker if expression of both your gene of interest and the marker are driven by the same promoter.
Here's some articles:
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I did some ribosomally depleted RNA-seq in rat brain. Does anyone have recommendations on how to quantify/compare small RNA expression between samples? I have used RNA STAR to map reads back to the genome and determine reads per gene for mRNA, but I can't seem to repeat that for any kind of small RNA like lncRNA or miRNA and the pipeline keeps failing. Is there a specific GTF file or similar type that I should be using? Do I need a different pipeline? Help please!
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Hi, Max!
In rRNA depleted libraries I believe you won't identify small RNA because probably the RNA fragments used to build the libraries were greater than you need to this.
However, to identify and quantify long noncoding RNA (lncRNA), you need a reference genome or transcriptome with this type of RNA. I've never worked with the rat genome, so I don't know what the annotation is like for that species.
The pipeline to quantify mRNA and lncRNA are the same, using the same reference genome. If you are not finding any lncRNA in the BAM file generated by STAR, it may be that this type of gene is not in your reference genome or that it has simply not been expressed in the condition you are studying.
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I did the experiment following the 2M sucrose cushion method to isolate the ribosome from 293F cells. Theoretically, I should get 10-15 A260 units from 2x 10^7 cells. But I can only get 4 units at most, I tried many ways, like increase the centrifuge speed, time, or lysis buffer, but neither of them works. Anyone with experience has good suggestions? Thanks.
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Particles smaller and less dense than the nucleus are obtained by gradually increasing acceleration on the supernatant. This operation is carried out on more powerful centrifuges such as high speed refrigerated centrifuges and ultracentrifuges. The order of illumination of the fractions is as follows: mitochondria, then membrane vesicles (vesicles) and ribosomes. The supernatant of the last centrifugation is "cytosol", i.e. soluble components of the cell that have passed into the buffer solution during tissue homogenization.
Further, by filtering, ribosomes can be isolated. But they will represent rRNA - supernatate
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Hi all,
I try to figure out weather there is a change in my rRNA following treatment.
when I treat the cells (mouse cells) and isolated RNA, I see a major difference in RNA concentration, without any difference in cell number.
what is the best way (or easy for start) to see if there is any change in the ribosomal RNA amount, activity or biosensis?
Thanks,
Shani
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Hi Shani,
I was just thinking about, if you already found proper primers for ITS and ETS in mouse, because I also want to quantify ribosomal RNA in mouse cells.
Am I right, that I have to use hexamer Primers für reverse transcription for quantifying rRNA. Oligo-dT-primers doesn`t make sense.
Thanks for your answer and help.
Katharina.
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Hello everyone,
I recently started several polysome profiling experiment and something bugs me out.
Every protocols I found use 12mL tube.
They load their lysate containing ribosomes attached to mRNA and then ultracentrifugate for 2h at 260000 g.
The more ribosomes attached to a given mRNA, the more it migrates. By analysing fraction of the gradient you can determine if your mRNA of interest is efficintly translated or not.
The tube contain a 10-50% sucrose gradient. This way, heavy complexes don't migrate to soon (and don'taccumulate at the bottom of the gradient) compared to light complexes.
I only have 4mL tubes instead of 12mL. So the lenght of migration is divided by 3. Hence, I also reduced time of centrifugation by 3.
But, is this "rule of thumb" accurate ?
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Agree with Pierre Béguin that scaling down will lead to loss in resolution of the polyribosome fractions.
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I have tagged my 12kDa eIF1 protein with Fluorescein-maleimide (Ex: 485nm, Em: 530nm). The interaction with this particular protein with 40S ribosome should show anisotropy value change. 10nM of eIF1_FL and 40nM of 40S in proper binding buffer (25mM HEPES-KOH pH 7.5, 2.5mM Mg(OAc)2, 80mM KOAc, 2mM DTT) were incubated and vertical and horizontal polarization measured. Anisotropy values calculated (Taking G=1) from those polarization values shows negative digits:
Blank Buffer: 0.123
Only eIF1_FL (10nM): -0.295
eIF1_FL (10nM) + 40S (40nM): -0.302
eIF1_FL (10nM) + 40S (200nM): -0.301
Please suggest how to improve my reaction conditions to get a values in the range of expected one.
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Because you did not apply a G-factor correction, your anisotropy measurements are relative. A negative value means that the perpendicular fluorescence intensity was higher than the parallel intensity.
Negative anisotropy is a possibility. The values can be between -0.2 and +0.4. However, in your case, it is probably due to the fact the the detection sensitivity of the perpendicular measurement is higher than the detection sensitivity of the parallel measurement. This is due to the instrument, not the sample. That is why a G-factor correction is used. In order to measure the G-factor, you require both horizontal and vertical excitation polarizer positions.
See this article for the definition of the G-factor and how it is applied.
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Hey fellow researchers,
one of the first things you learn as biologist is the structure of the cell. So basically, every biologist should be skilled in the basic terms. But I encounter very often (even in very high ranking journals, including Nature, Science and Cell) the mix-up of cytoplasm and cytosol.
The definitions, as far as I know, are very clear.
The cytoplasm is the complete content of the cell surrounded by the plasma membrane, which includes the cytosol and ALL membran-enclosed organelles (with excpetion of the nucleus), as well as solutes, the cytoskeleton with all components, ribosomes and so on.
The cytosol is the fluid phase of the cytoplasm without the membrane-enclosed organelles.
So the cytoplasm includes the cytosol but not vice versa. Therefore, sentences like "Protein X was detected in the cytoplasm, but not in ER and mitochondria" is wrong, since ER and mitochondria are part of the cytoplasm and one cannot exclude them when using the term cytoplasm as definition for a cellular location. And this is only one example of many.
Do I miss something or have the terms changed? Perhaps experts from the field of cell biology can contribute to this discussion and help to understand.
All the best and stay healthy,
Marc
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Hey Jasper and Suraiya,
thank you very much for your responses. Of course, if you seperate the organelles, but keep the rest, this rest still will bre the cytoplasm just without the organelles removed. Absolutely right.
But I got another example (describing the whole cell situation): "Mitochondria realease cytochrome c into the cytoplasm". It is not wrong, but it suggests, since mitochondria are part of the cytoplasm, that cytochrome c is also released into mitochondria (the matrix for example), which in this case is not the point the authors want to make. They just mistook cytosolic release of cytochrome c for cytoplasmic release. What do you think? Looking very forward to hear from you.
All the best and stay healthy :)
Marc
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Does anyone know if eukaryotic ribosome translation / integrity can be preserved in other buffers than HEPES and TRIS after purification through a sucrose cushion, by IP, or other methods?
Maybe a reference where Ribosome activity or integrity in various buffers (and not only Mg2+ and K+ concentrations) is tested? Could be in vitro translation, cryo-EM, or other methods.
All the best - with hopes for any helping thoughts,
Kasper
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Nice Contribution David Farringdon Spencer
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Hi,
I have stumbled upon an article about translocation of AKT into mitochondrial matrix upon IGF stimulation and as I tried to find some mechanism of this transport I found another articles about translocation of cytosolic proteins into mitochondria (GSK upon EPO treatment, HSP60 upon dehydratation).
I have always been taught in our cell biology lectures, that nuclear-encoded proteins are imported into mitochondrial matrix predominantly co-translationally and that only small portion of proteins are translated on free ribosomes but are kept unfolded by chaperones and then, still unfolded, transported into mitochondria, where they acquire native conformation.
Since this is not main focus of my work, but I find it interesting, I want to ask if somebody knows how well studied and how common this translocation of mature cytosolic proteins into mitochondria is and weather other such well-known proteins do this. And maybe suggest some review/article on this topic.
Thank you very much
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Thank you very much for our answers. I forgot to mention that I´m only interested in human cells. The TAT machinery seems not to be conserved in humans, so i guess its not the thing I´m looking for.
I´m also aware of te existence of pathways transporting unfolded precursor proteins into mitochndria. What I´am actually interested in is wheter there is some known pathaway that folded, functional cytosolic proteins use to translocate into (and possibly out of) mitochondria upon certain stimuli.
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Hello everyone, I have a question about the translation rate. I want to check the translation rate of isoforms belonging to one specific gene. For example, isoform A and isoform B of X gene, which one goes ribosome more and the translation rate is high. Can you give advice about technique and research paper? Thank you :)
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translation process need amino acid add for elongation of polypeptide chain
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The answer choice is C.
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Hello everyone, I recently came across something very peculiar called internal ribosome entry sites (IRES). These secondary structures are present at the 5' end of many viral as well as mamallian mRNAs and are required for cap-independent translation (ex- at the time of apoptosis). Can somebody explain how the mRNAs are folded into such complex secondary structures? Are there any factors involved that regulate this folding or the mRNA itself catalyses this reaction?
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I don't know about this, but here is a review of the subject:
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Experiment sample : Rhizobacteria.
After screening and isolation and the extracted of bacterial DNA by using 16s ribosomal technique to get sequencing result (in the form of nucleotide sequence).
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you can use the ediseq and finTV exe to anylize the result
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Hi,
I want to separate 60S from 80S ribosomes on a polysome gradient. I usually run 10-50% sucrose (3h at 35k rpm), but then they bleed into one another and the resolution is not good enough to tell if my protein binds to 60S only or to both 60S and 80S. I have also tried 10-25% (4h at 35k rpm)and 10-30% (3h at 35k rpm), but they either completely get rid of the 80S or they still keep the 60S and 80S together.
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Hello Kamena,
You can check the Basic Protocol 2 from Analysis of Eukaryotic Translation in Purified and Semipurified Systems by William C. Merrick et al., Current Protocol.
'For analysis of 40S to 80S complexes in 10% to 25% sucrose gradients, centrifugation is 16 hr at 55,000 × g (20,000 rpm in SW28.1 rotor), 4°C. This completely pellets the polysomes while separating the 40S, 60S, and 80S complexes into distinct fractions (see Fig. 11.9.3). By increasing the sedimentation speed of these gradients to 23,000 rpm, 80S complexes are also pelleted, and separation of 43S from 48S (preinitiation) complexes is possible.'
Hope this protocol help!
Good luck on your experiments!
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Hi everyone,
I was wondering what antibodies or pcr primers I would have to use to control my fractions in a polysome profiling experiment ?
A quick search highligthed antibodies against small and large ribosomal subunit like rpl17a and rps3. And anti-Ssb1 for soluble fraction ?
Can you me give me advice on this topic please ?
Thanks,
Philippe.
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Hi Philippe,
For western, you can use rpl17a and rps3. For PCR, we generally use any housekeeping genes such as gapdh, actin, hsp 90 for normalization.
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Is it possible for ribosomes to recognize mRNA and initiate translation without the shine-dalgarno sequence and initiation codon? In the Nirenberg -Matthei experiment with poly U mRNA and E.coli cell free systems that helped in cracking the genetic code, there is no mention about these requirements. My doubt is that how was it possible to initiate translation in a poly U mRNA (or any other synthetic mRNAs) without the recognition and initiation signals?
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Thank you Mr. Levine for your response.
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I want to targeted MD simulation of macromolecule using AMBER software .As i am quite new to MD simulation field and of such a big complex .
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Thank you.I will try using that
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I intended to perform polysome sequencing upon FAC-sorted C. elegans cells, but the required number of cells is not achievable for my application. Ribosome nascent-chain complex (RNC) sequencing seems like a viable alternative for which fewer cells are needed - but does anybody have an estimate of how much the minimum requirement is?
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I use 4 concatenated molecular markers for phylogenetic inference. One of them, ribosomal 16s, shows a large amount of indels in my group of study, and I would like to use that information which is otherwise lost.
Could you point me to a thorough description of the best procedure for coding gaps as an additional partition and incorporating them in RAXml/MrBayes (substitution model, etc.)?
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FastGap is a Windows executable program for fast and efficient assembly of DNA sequence alignment files. In the process, gap or indel characters can be coded and added to the data file as separate partitions:
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Ribosomes are described as ribonucleoprotein structural complexes with the definite function to direct protein synthesis. If the structure that is supposed to make proteins already has proteins as part of its composition, where were those proteins made giving that the protein components of ribosomes pre-date the protein-synthesizing apparatus?
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As already mentioned above, ribozymes are thought to have mediated protein synthesis in early days.
Here is the first "proof of concept" paper as far as I know.
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I am using Trizol to do RNA extraction from peritoneal tissues (rat). The samples were stored in RNA later. I followed the Trizol protocol exactly but i get two (18 and 28s) for some samples and only 1 band for others. The RNA extraction for the samples were done on the same day and at the same time. Can anyone please help.
Thank you.
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Thank you for your help Laurence.
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The En3hancer spray is no longer sold. Does anyone have a protocol for performing 3H-pulse chase analysis of yeast rRNA where the nylon (no the gel) is subjected to other tritium enhancer ?
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They are reusable !
but pay attention that your nylon is completely dray before screen exposition
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Hi everybody!
The whidely used method to quantify the various ribosome fractions (40S, 60S, 80S and polysomes) are by sucrose gradient and fractioning.
Unfortunately this techniques uses very high amount of cells (more than 10 million or even more).
I need to make a knockdown in a suspension cell line of a gene throth siRNA and than to evaluate the relative 80S/40S/60S ratio (not interested in polysomes) but It is difficult to knockdown such high amount of cells.
Does anybody know another technique to quantify the ribosome fractions starting with a lower cell quantity?
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Hi Roberto,
Option Nr. 1 would be to use a different (more long lasting or stable) approach to get rid of your gene of interest, something like CRISPR. This would then allow you to grow as many cells as you need for polysome profiling. However, there are probably good reasons why you don't want to do that.
Option Nr. 2 would be the following, but I have to say that I have never tried it myself: Next to high quality input lysates, the most critical part of a sucessful polysome profiling experiment is the choice of 'steepness' your sucrose gradient (i.e. 20%-70% versus for example 60%-80% or 20%-50%). Such different gradients allow you to have a different dynamic range and resolution of your fractions. If you want to purify mainly polysomes you will need a different gradient compared to when you want to purify only the small ribosomal subunit. Since you are not interested in polysomes, but the ratios of 80S/40S/60S you will need a gradient that resolves well in the "low" molecular weight range, for example 20%-50% sucrose. That way, all polysomes that you are not interested in, will be stuck in the last fraction and they don't annoy you. But maybe you knew all this already. The important point is this: For your exact cell type you will have to find the best gradient. Therefore, I would propose to perform this experiment with a sufficient amount of you cells (but importantly no knockdown) and find the best gradient for you. Try to aim for as much 40S, 60S and 80S in single, but different fractions as possible.When you have found a good gradient for your cell type and biological question (high resolution of 80S/40S/60S ), I would suggest to use exactly this gradient for a smaller amount of your knock-down cells (use as much as you can possibly get without spending your entire budget on siRNAs...). Depending on the sensitivity of your hardware (absorbance measurement etc.) and final amount of cells, you might of course not see peaks. However, from your earlier characterization of the non-KD cells you already know exactly which fraction contains the 40S, which one the 60S and which one the 80S. To visualize this information and to semi-quantitativly analyze the ratios, you should then perform a Western blot against exclusive 40S and 60S marker proteins. You could use for example an antibody against RPS27 (MW: 9.5 kDa) to detect the 40S and an antibody against RPL9 (MW: 22 kDa) to detect the 60S. The 80S will of course contain both proteins. A Western blot containing all your fractions probed against both 40S- and 60S-specific antibodies should then give you the answer how the 80S/40S/60S ration changes with and without knock-down (re-do the experiment also with the exact same amount of cells for the non-knock-down condition to better compare the relative ratios).
Important: If you want to isolate RNA for sequencing this will not work, since quantities will be too low, but for Western this should not be a problem due to the high sensitivity.
I hope this makes sense. Good luck!
Johannes
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Enzymes are synthesised as zymogens and then make a transition to active form when required, is this true for every enzyme or are there any exceptions to that?
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Those synthesized as zymogens are actually rarer than those that are not. For example, almost all enzymes synthesized for major metabolic pathways such as glycolysis and the TCA cycle do not require activation from a zymogen form.
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Protein biosynthesis in an RNA world could have had the following components: ribozymes instead of ribosomes, flexizymes instead of aminoacyl-tRNA-synthetases, mRNAs, tRNAs, other kinds of RNAs. Is there anything more needed for protein biosynthesis? What?
(Does the environment of early genetic code evolution have enough ATP? Are proteins needed for proper tRNA folding?)
Are proteins needed for protein biosynthesis or is RNA in the right environment enough to build a translation process?
Thanks in advance.
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Matthias R. Schaefer thanks for your interest. My question is about the end of the RNA world and the roots of protein biosynthesis. I want to know how the transition could have been. There are lots of theories but I found none that consider many aspects. There are theories about the first amino acids but not what the translation process they took part in looked like and theories about aaRSs withouth codon recognition domains but where did these aaRSs come from when there was no translation process before their existence?
I appreciate any piece of information related to the topic no matter if it's knowledge, theory or hypothesis or a couple of reasonable assumptions.
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The EM image of E.coli mRNA being transcribed by polysomes by Miller et al. in 1970 became iconic The sample preparation, however, seems particularly involved.
I want to know what options I have today if I want to image transcription (from a plasmid) and/or translation in E. coli?
Thank you very much
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There is a great, molecularly correct, movie of translation in this movie called:
The Molecular Basis of Life
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Title explains itself. I am trying to find information about a question in my mind.
For example if we divide a cell by regions like west east noth south, a tRNA going out to look for its spesific aminoacid, it advence in cytosol(for example, to the east from the west) and find the a.a it interacts(at east region of the cell), take it to the ribosome(for example at north of the cell) and take off.
But, how?
How tRNA knows that a.a is at east and how it knows the way to ribosome after binding a.a?
is there something like electrostatic forces made it to advence in correct direction? and cytosol act as conductor?
Could you give me articles about this issue, I could not find a study other than interactome studies but interactome studies is not the answer i am looking for. Interactomes only answers if molecules are interaction close. But I want to know how distant molecules find each other?
Thanks and regards....
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Simplistically, molecules diffuse through the cytoplasm because of their random motion. The rate of diffusion is affected by the viscosity of the cytoplasm and by the size and shape of the molecule. Since the dimensions of most cells are quite small, it does not take long for freely diffusing molecules to encounter their binding partners. Specialized transport systems exist to transport molecules over long distances, such as in the axons of neurons.
You should look for literature on the diffusion coefficients of biological molecules in cells. Here are a few articles:
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I read an article where they said, they used 20-25 OD A254 sample for a ribosome separation on sucrose gradient. I know this is an important parameter but I am not getting it how much does that mean? Can anyone explain it please?
Thanks
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The OD unit is a typical measurement of DNA or RNA, with 1 OD260 unit equal to 40 ng/µI of RNA, for example. What is missing from this definition is the pathlength through the sample. It refers to a 1-cm pathlength, the usual pathlength of a quartz spectrophotometer cuvette:
I think the authors meant that they used a quantity of ribosomes that would have had an absorbance of 20-25 if measured in a 1-cm pathlength cuvette at 254 nm in a 1 ml volume.
Of course, it is not possible to measure such a high absorbance. A small portion of the sample would have been diluted for the absorbance measurement.
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Hello folks,
I'm interested in looking at ribosomes by negative-staining, and I understand that a lot of people use the FEI Morgagni for this purpose. However, the only TEM that I have access to is a Jeol 2000FXII. Is there anyone here that has used this EM for looking at ribosomes (or other macromolecular complexes?)
Best wishes,
Pramod
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Hi Pramod,
As Dr. Gowen said, any TEM is Ok. For negative staining the resolution is limited with sample preparation, not with microscope's properties. It would be better to use pure carbon film grids, not formvar. But
The best way is to use a good cryo-TEM like FEI Titan Krios.
Good luck!
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Dear sir:
Regards
Last year, I've studied the Tet resistance among Aeromonas spp. and I found that all tetracycline-resistant isolates carried at least one of the tet genes examined. Among efflux genes, tet (A) was most commonly observed in isolates, followed by efflux tet gene (tet B and C) and ribosomal protection protein (tet O). However, efflux genes tet (D and G) were not detected in any of the isolates.
look at the link below:
http://www.bldeujournalhs.in/article.asp?issn=2468-838X; year= 2017; volume= 2;issue=1;spage=22;epage=28;aulast=Al-Charrakh
Regards
Prof. Alaa Al-Charrakh
Babylon Univ., Iraq.
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I am in an academic research project, where we want to introduce a gene that codes for obtaining an enzyme that degrades phenols in the organism P. Infestans, so we want to introduce a construct together with the enzyme of interest in the organism, and this It leads me to ask myself the following question: We want to introduce the gene near the ribosomal site since it is a very conserved region and therefore this would prevent the gene from being maintained in the organism, so could this be possible? Or is it better? place the construct in another part of the body's genome?
Greetings.
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آسف السؤال ليس من اختصاصي
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What is pathophysiological Significance of mutations in the Ribosomal DNA (rDNA)?
Hence in human molecular genetics, we focus on mutations/variants in the genes encoding mRNA and then proteins to predict their possible impact in genetic disorders.As rbosomes are assemblies of proteins and rRNA molecules that bind to and translate mRNA molecules to produce proteins.
1. If there is a mutation/variation in these genes encoding rRNA what would be their possible impact?
2. If the sequence of the transcribed rRNA is disturbed, how can it disrupt the ribosomal assembly i.e. binding of rRNA, ribosomes and mRNA?
3. If this assembly is staggered, then the transnational process might be definitely affected?
4. Is there any genetic disease reported with mutations in the genes for rRNA?
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Irfan, Please let me know if you find the attached papers informative to the underlying basis of your question. This is an area of investigation that we are actively pursuing. I think our most recent papers on the subject open up the discussion a bit further than is afforded by the functional/non-functional paradigm.
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Membrane proteins can be inserted into membranes via different mechanisms and in different orientations. I am specifically interested in mammalian cells (not bacteria or yeast). Does anyone know how high the percentage of type I, type II, and type III proteins is among mammalian integral plasma membrane proteins? Is there a review out there that has some numbers?
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Hi Ralf Max Leonhardt sorry, I cant find any specific references on the abundance of membrane proteins, but I gather that most type-III membrane proteins are housekeeping proteins like ion pumps.
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I am removing enzymes, amino acids and other small molecules from ribosome complexes through density gradient centrifugation with sucrose. I have very little ribosome sample so I would behave with volume as small as possible to recycle my samples. Could anybody tell me what's the minimum volume of an efficient density gradient centrifugation? 100ul worked for me, but I am not sure if I behave 50ul still works without losing my ribosome.
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  1. Centrifugationhttp://www.phys.sinica.edu.tw/TIGP-NANO/Course/2007_Spring/Class%20Notes/AC_Chapter%203%20Centrifugation%200321.pdf-Nucleic acids. Basis of separation: -Size. -Shape. -Density. Methodology: - Utilizes density difference ... 13. Relative centrifugal force. RCF = 1.12 x 10-5 x ( rpm)2 x r r min r max .... Two different types of density gradient centrifugation
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I am basically interested in understanding the graph obtained after ribosome profiling and how can you know from the graph in which translational state are the mRNAs?
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I would rather like to extend a question in this discussion. I want to check translational block of a gene using polysome analysis but I m confused how to differentiate between actively translating genes from translationally blocked genes??
because both kind of genes could have multiple ribosome units on them. Kindly shed some light.
Many thanks..
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Hi, dear colleagues,
Recently, I really confused about ORF in plasmid. I checked my target sequence in my plasmid, in fact, after the promoter there are two possible ORFs, 1st ORF is close to my promoter, as a 56aa peptide, 2nd ORF is my target protein 219aa [CmR].
In my opinion, if the 1st ORF is translated, the 2nd ORF will keep silent and no protein production. The fact is that the 2nd ORF is translated well, the 1st ORF is silent, even it is closer to promoter. The sequence file is attached.
My questions,
Whether there is a limit for ORF length?
>In SnapGene default parameter, the ORF should bigger than 75aa, but I’m sure the short peptide could be translated. Maybe in plasmid not ??
Always the first start codon [M] could be recognized during the translation, I wonder whether the small ribosomal could choose the start codon that is included in a longer ORF. That’s really incredible, if so.
Thanks for your any help~
Yunlong
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Thanks for your answers, Pietro Pilo Boyl and Yazan J Meqbil !
This sequence is retrieved from a commercial plasmid [#addgene], so I trust it as right and reasonable construction.
I am trying to understand why it works. As you said, the target gene [CmR] is translated thanks to SD sequence. It is important to recruit the ribosome to the messenger RNA (mRNA) to initiate protein synthesis. Besides, an mRNA codes for multiple polypeptides in bacteria is possible. Maybe these two ORFs could be translated. One with SD sequence is highly-translated, the other w/o SD is poorly translated.
Then I checked the sequence of CmR, yes, surprisingly, Sequence is started with a good Kozak consensus. I found explanation about it, " Eukaryotic ribosomes (such as those found in retic lysate) can efficiently use either the Shine-Dalgarno or the Kozak ribosomal binding sites. " [from https://www.thermofisher.com/fr/fr/home/references/ambion-tech-support/translation-systems/general-articles/ribosomal-binding-site-sequence-requirements.html]. It seems that SD+Kozak+ORF in palsmid is to ensure its wanted translation.
However, could I say that the SD is necessary for a prokaryotic translation plasmid ? Because it allows the bacterial ribosome to be built at an interior position on the mRNA through direct binding to this sequence.
And Kozak sequence is not necessary in an eukaryotic ORF in plasmid ? Because I found many ORFs in our plasmids express well, without Kozak sequence. Or maybe, if a strong promoter , needn't Kozak ?
Thanks again~
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I am currently using a lac-based vector for periplasmic expression of proteins in E. coli. I now want to re-design the vector and express the proteins under the control of the T7 promoter in a DE3 strain. I found a lot of different vector sequences and am a bit confused which sequence parts I really need in my vector for IPTG- inducible expression of my proteins.
T7 promoter only? T7 promoter and T7 terminator? T7 promoter plus lac operator? (If I got that right, the T7 RNAP is under the control of a lac promoter in the bactrial genome, so I shouldn't need a lac operator in my vector?) Which ribosome binding motif?
I am looking forward to your replies!
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If you want to create an IPTG-inducible system in E coli, you actually have two choices: 1) you can have the lacIq repressor module encoded within the vector itself, as is the case with the commonly-used pQE-80L backbone (https://www.addgene.org/vector-database/3883/). In such a case, you put your gene of interest in front of the T5 promoter and could express it in an E. Coli strain like DH5-alphas (which are typically used for MiniPreps, rather than expression). 2) Your other choice is to use an expression-optimized backbone, such as pRSET and its variants (https://www.thermofisher.com/order/catalog/product/V35120) and place your gene of interest in front of the T7 promoter. You then transform this into the popular E. coli strain of BL21(DE3), where the λDE3 prophage was inserted in the chromosome of BL21 and contains the T7 RNAP gene under the lacUV5 promoter. This strain and its derivatives is by far the most common for protein expression. In both cases above, you may wish to add some dextrose solution to the culture as it grows to inhibit protein production until you reach the proper optical density for protein production (between 0.6-1.0) before then adding your IPTG to induce expression. I hope that helps!
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If I mutate a start codon, how far along a transcript will a ribosome scan for a downstream AUG before it gives up and falls off?
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This is an interesting question.
ATG is not the only recognised start codon. GTG is almost as good and CTG and TTG are also used.
The next part to be considered is the Kozac sequence preceeding the start codon. A strong Kozac can over come a poor start: AAA TTG may work
Less important but relevant is the presence/absence of secondary RNA structures. These can increase or decrease translation.
At the moment prediction tools are inadequate to make reliable statments about the effect of mutation of start codons. Experiment is the best way to find out - at least at the moment
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Hi all,i'm currently studying a brain sc-seq data by seurat package,and my cluster analysis seems to be profoundly influenced by genes that relate to ribosome(named Rpxxx).A few clusters is characterized by expression of these genes,and known markers of cell types seems to have little relation with these clusters.I have regressed cell cycle by ScaleData function,so i'm confused about what these ribosome-related dada means.Shall i delete these data?Or shall i study these ribosome-related clusters in other way?Thank all!
BTW,i have deleted blood cells from my data by hemoglobin-related genes
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Hi,
I tend to be quite careful about removing genes in the data set if I see that the data quality is good. Have you performed cell quality control (filtering out cells based on counts/number of genes/mitochondrial RNA fraction) and adequate normalization and batch correction if necessary? I have also previously found clusters with a lot of ribosomal gene markers, but looking at known marker gene expression clarified this (in my case Stem cell markers).
Also, is your data set mouse or human? If it is mouse you could check out Linnarson's mouse brain atlas.
Good luck!
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Hi, I am new to this. I have done sequence alignment of modified tRNA genome sequences from E. coli and Mycobacterium tuberculosis against the whole genome database of tRNA and have not found any hit showing the similarity in their sequences. I am trying to do Mycobacterial translational complex studies and would like to know how relevant it is to reconstitute Mtb ribosome with E. coli tRNAs. My query protein binds to tRNA as well as mycobacterial ribosome in vivo. I am performing biochemical and structural studies.
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I am not speaking from direct experience, but considering the lack of sequence similarity between M. tb. and E. coli tRNAs you mentioned, I would not want to use E. coli tRNA for the study. It is probably quite difficult to prepare tRNA pools from M. tb, however, so you may instead choose to use M. smegmatis. At least it's in the same genus, and it is easier and safer to work with.
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I am unsure what the differences between IRES1 and IRES2 are at the sequence level, whether IRES2 is a development of IRES1, and what the consequences are of any modifications to the original sequence.
I'd be very grateful for any pointers or literature references.
Thanks a lot,
Jonathan
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An IRES initiates translation in a cap-independent manner, allowing synthesis of two proteins from a single bicistronic mRNA The source of this sentence is from the site listed below, which does a very good job of explaining these RNA sequences.
Hope it helps.
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Depending on the answers of my first question, I would also like to know how to get rid off these ribosomes from the mRNA after doing IP of a particular protein. In precise, I want the condition where I can only get the particular protein bound to translating mRNA.
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Thank you so much Tony.
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Why all the amino acids that are ribosomically incorporated into proteins exhibits L-configuration not D-configuration?
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Hi Hirakjyoti,
My immediate guess is since amino acids are incorporated into the proteins in the L-form during the translation, it follows that they are also metabolized and biochemically processed in the L form. It is possible that the translation process dictates the L-stereoisomer amino acid to be the isomer which evolved and functional in the human body, involved in metabolic processes.
Best,
Hediye.
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All enzymes undergoes post-translational modification (PTM) to become a functional mature enzyme. Is there any enzyme which doesn't undergo PTM? any exceptional enzyme?
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I do not think it is a general rule that all enzymes undergo post-translational modification. It is definitely not true of many (perhaps most) cytoplasmic bacterial enzymes. I know this because the measured masses of many bacterial proteins expressed in E. coli match the predicted masses precisely. (Caveat: reversible covalent modification such as lysine carboxylation may not be observed by this technique.)
There is surely a lot more PTM going on in eukaryotic cells, but I doubt that every cytolasmic enzyme is modified. There may also be cases where a protein is sometimes modified in an accidental way (e.g. phosphorylation) that has no significant bearing on its function.
If you disregard removal of signal peptides and formation of disulfide bonds, then you can probably include some noncytoplasmic enzymes among those that undergo no functional PTM.
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Just wonder is there any evidence exist between ribosome copies and number (or diversity ) of the proteins as end product in a cell?
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