Ribosomes - Science topic

I guess NO!

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.

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.

Hi, Maria

4V6I is a ribosome with an mRNA, for example.

Regards,

Alexander

Yes it works, because fish belong eukaryotes. IRES has role in translation process.

Hi Matt Wu ! Yes, we did and saw the same thing. Please find attached the bioanalyzer results. The RIN is 8.7.

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.

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

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 :)

Did your experiment include an IgG control or similar? This would help with interpretation of this result. Definitely an odd one!

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).

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.

Aashish Bhatt I actually want to do AA only in this small subshell.

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.

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

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.

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

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.

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.

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.

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.

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).

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.

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.

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!

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.

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.

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:

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.

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

Regards, Sergey Viktorovich Pushkin

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.

Agree with Pierre Béguin that scaling down will lead to loss in resolution of the polyribosome fractions.

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.

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

Nice Contribution David Farringdon Spencer

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.

thank you Enes Yağız Akdaş and Enes Yağız Akdaş

The answer choice is C.

I don't know about this, but here is a review of the subject:

you can use the ediseq and finTV exe to anylize the result

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!

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.

Thank you Mr. Levine for your response.

Thank you.I will try using that

I hope this helps:

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:

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.

Thank you for your help Laurence.

They are reusable !

but pay attention that your nylon is completely dray before screen exposition

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

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.

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.

There is a great, molecularly correct, movie of translation in this movie called:

The Molecular Basis of Life

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:

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.

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!

آسف السؤال ليس من اختصاصي

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.

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.

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..

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~

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!

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

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!

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.

Here are a few articles on the mechanical properties of cells.

Regards,

Joachim

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.

Thank you so much Tony.

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.

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.