Questions related to Protein Expression
I am new to the world of protein expression and purification and have no prior knowledge or experience about this topic. I will be starting to express my first 2 proteins in a couple of weeks. It will be the gpW and engrailed protein to be expressed using E. Coli. system. Before I start, I would like to know what background knowledge I would need before getting started (I have majored in chemistry with an emphasis on physical chemistry). Can you point out some resources from where I can build my knowledge, and get tips on the procedure and troubleshooting?
I am trying to express several proteins at the same time, but I want to use a different promoter and terminator for each one to avoid the possibility of recombination.
The promoters that are available to me are: TEF2, PGK1, CCW2, TDH3 and HHF2. The available terminators are: ENO1, SSA1, ADH1, PGK1 and ENO2.
Has anyone ever used these combinations of promoters and terminators? In your experience, which combinations work the best?
I do not like how my Keyence BZ-X710 images have been coming out lately. I’ve been noticing grid-like shadows on the images after stitching them, but not while looking at the live images. The Keyence halide bulb was the original culprit, but the problem is still apparent despite it being replaced. I care about this because I’m looking to quantify protein expression, which feels pointless if the image brightness and contrast looks messed up. Slices are 40 microns and are stained for IBA1, GLT1, and DAPI. They’ve only been imaged 1-2 times.
What do people think could be causing this? How do you recommend I go about fixing this?
Anything help. Thanks in advance!
Hello. I have a problem. I am expressing a protein in the SoluBL21 strain at two temperatures (18°C and 20°C). At 18°C the pellet was beige while at 20°C it was gray. Generally, in other cultures that I have done with the same bacteria, it has not looked as dark. What could have happened?
Hello. Hopefully, everyone is doing well. I usually use the Pichia protein expression system to express my recombinant protein. Every time working with this system was so easy for me and had no problem at all but for the last two months whenever I try to express the same proteins in this system, after purification i notice lots of DNA contamination that cannot be removed from my protein sample. I want to know if anyone faced the same problem before as I do not understand what has been changed in my procedures which make me face this huge problem. If anything you can mention which may help me is really appreciated. (I also tried so many wayed to get rid of this contaminant but was not useful) THANKS
I did a knockdown, after checking the level of protein expression with Western blot, I got some percentage let's say like 60 to 70 % of knockdown, but when i tried checking with qPCR to check the transcription level it was as if there was a complete knockout, what could have been the reasons, I am confused can some help me with an explanation? You help will be highly appreciated.
I'd like to know what's current progress of 'synthetic circadian clock' or 'synthetic oscillators'. Indeed, I would like to investigate methods to regulate protein expression (e.g., transcritpion) in a time controlled manner. Being umfamiliar with this field now, I am eager to know what are already known, and what current designs are effective?
If you know papers of importance or groups with expertise, please note their name.
Thanks for your time.
I'd like to perform a SEC step prior to on-column refolding of a protein I am expressing/purifying but am worried about the extended time the protein will be in the presence of urea (and subsequent protein carbamylation) in the current refolding workflow I have.
Is there any concern of protein modification with a 4-6 hour denatured protein purification workflow using 6M GndHCl?
I have a synonymous variant library of a protein, and it has hundreds of variants. They are cloned in the Flp-In T-REx expression vector to work with the Flp-In system. We have worked with this library to measure the protein levels using the Flp-In HEK293 cell line and it has always worked. Right now I would like to transfect this library into other human cell lines and unfortunately, these new cells do not have the Flp-In T-REx landing pad and it would require a lot of work to generate them.
I wanted to ask if there is any other high throughput method to measure the protein levels of these variants in human cells.
Thanks a lot.
I am trying to express a novel GlcNAcT enzyme in BL21(DE3). I have successfully cloned it in pET28a vector and confirmed it through sequencing. I tried following previous protocols for GlcNAcT expression but did not see any protein expression. Induced and uninduced samples both are looking the same in SDS-PAGE. What might be the cause, and how can I resolve this situation?
I am interested in the relationship between gene dosage and the amount of protein expression. Any one has experience in this concern? If there is a consensus?
I am trying to produce an antibody from a plasmid in Freestyle CHOS cells. The plasmid is a HC/furin/p2A/LC vector. The cells were transiently transfected during the exponential phase and we see expression after purification (protein A) and intracellularly. The issue is, I mostly see LCs and not much HCs. For some reason, the intracellular bands look like attached. The bands were stained with a secondary antibody targeting human LC+HC. There's no band at 50kDa, but a double band around 40 kDa. The LC is stained at 25 kDa.
And native protein electrophoresis showed a low amount of HC and assembled IgG, and predominantly LCs.
Does anyone know what the intracellular 40kDa band should be? And since LCs themselves should not even bind to protein A tightly, is it reasonable to have eluted majorly LCs, and not much HCs? Finally if anyone has suggestions on how to improve HC folding and full length IgG assembly, that would be helpful. Thanks in advance.
I would like to know if someone already used the pRSET plasmid into normal BL21(DE3) and not into BL21(DE3)pLysS for protein expression.
I'm having some induction and purification issues and i wonder if the problems could come from the utilisation of normal BL21(DE3).
I have been trying for months now to get my protein expression confirmed by western blot and failed every single time (I checked the antobodies, they are working fine). I have transformed yeast (Pichia pastoris KM71H) with my plasmid by eletrocporation (4 kb long, my protein is 26 kDa, 1210 bp) confirmed transformants via PCR but I don't get any protein produced afterwards. Does anyone have any idea on what is going on? Is it possible that my confirmation on PCR is actually a false positive result?
If someone could help me with that it would be awesome, thank you in advance!
- I am working on Immunohistochemistry for Per1 protein expression on mouse brain coronal sections (40microns thickness, free floating).
- In the protocol that I'm following, it says that the DAB exposure time is 1-30mins and I have observed different people using different exposure time.
- 30mins can cause too much background staining and 5mins barely did anything for my protein staining.
I have an Arabidopsis protein, which should be apoplastic (rice homologue is localized to apoplast and this protein contains predicted signal peptide).
I expressed the protein without the signal peptide (but I have also version with the SP ready) with the alpha factor under control of AOX1 promotor. After dilution of O/N preculture, I expressed the yeast (also plasmid control, which should express protein in frame with the C-term tags) in the BMXY medium, which I supplemented with half of glycerol and half with methanol. Next 4 days, I supplemented methanol to keep the induction and I collected samples every 24 hrs, when I centrifuged 1 ml of medium and froze the supernatant and pellet (cells) separately.
As I did not detect activity in the supernatant previously ( https://www.researchgate.net/post/Can_a_plant_cell_wall-associated_protein_remain_in_the_cell_wall_when_heterologously_expressed_in_yeast_Pichia_pastoris ), I extracted proteins from the cells and from cell wall and loaded them on SDS-PAGE. I wanted to load the same amount of proteins to each well, so I loaded rather low amount, because medium contained only low concentration.
I have used an old polyclonal anti c-Myc antibody, but there was nothing on the blot, except of some non-specific bands in the control.
As there is no clear band neither on SDS-PAGE, nor on Western blot, I would consider this as negative results. So what can I try next?
First, I could try gel and blot with more proteins. However, since there is no obvious large band, does it make sense?
Second, I could check all timepoints for expressed protein.
Third, mRNA analysis.
What should I do next? What makes sense and what doesn't?
We have constructed several plasmids that express HA- tagged proteins. At first, all the protein expression of these plasmids were confirmed by WB. But now, after several rounds of amplification of these plasmids, the expression of HA tagged proteins could not be detected. Does anyone have this problem?
I have used western blot to check SIRT1 and H3 acetylation expression. I saw change in protein expression of H3 acetylation but no change was seen in the SIRT1 expression. I got same intensity bands across all the samples. Why is that?
Can I culture HEK293T cells and MSCs together (co-culture)? What would be the best method?
How can I extract RNA from HEK293T only to monitor gene expression from being co-cultured with MSCs (simply put, I want to monitor the effect of MSC through gene/protein expression)
I am experiencing issues with the expression of phosphorylated proteins in my western blot experiments. Specifically, I observe strong phosphorylated protein expression but no expression of the corresponding total proteins in the same samples. For example, I detect phosphorylated STAT1 (pSTAT1) but not total STAT1 protein. Similar results were obtained for pSTAT3 and STAT3. I have thoroughly searched online but have been unable to find a possible explanation for this phenomenon.
I would greatly appreciate any advice or suggestions from anyone who has encountered a similar issue.
Here is my experimental protocol:
- Prepared single cell suspensions from fresh mouse spleens using a buffer containing 1x PBS, 2% FBS, EDTA, and antibiotics.
- Washed the cells once with ice-cold PBS and then lysed them using RIPA buffer (with proteinase and phosphatase inhibitors) by vortexing for 10 seconds every 5 minutes on ice, repeated 4 times.
- Quantified the protein concentration using the BCA assay and mixed 30 micrograms of protein with loading dye, boiling the mixture at 90℃ for 10 minutes.
- Transferred the proteins to membranes and blocked the membranes with BSA at room temperature for one hour on a shaker.
- Washed the membranes three times with TBST containing 0.2% Tween-20.
- Incubated the membranes with primary antibodies overnight at 4℃ on a shaker.
- Washed the membranes three times with TBST containing 0.2% Tween-20.
- Incubated the membranes with secondary antibodies at room temperature on a shaker.
- Washed the membranes three times with TBST containing 0.2% Tween-20.
- After detecting phosphorylated proteins, stripped the membranes by adding deionized water and microwaving for four minutes.
- Blocked the membranes with BSA and incubated them with primary antibodies.
I am finding it hard to understand the difference between protein expression and what is protein activity. Like if my protein is getting expressed in western blot how can I relate it with its activity? I used ELISA kit that tells me about the concentration of SIRT1 in samples. But I wanted to know whether the activity of SIRT1 is being inhibited or activated. How can I know that ? Do I have to use some other kit?
I have cloned a mammalian gene through the gateway cloning (Phase Lamda ) method. The codon optimization was done via IDT tools. The orientation of the insert has been confirmed by Sanger sequencing.
The vector was transformed in E coli BL21 AI cells. recombinant protein expression conditions are :
1. inducing protein expression with 0.2% L-Arabinose, when OD reached 0.5-0.7.
2. Expression temp 37 for 4 hr.
Thankss in advance
Recently in one of our tests we identified a cellular protein expressed without any mRNA expression of the same protein intracellularly. What could be possible explanation?
Kindly add possible references.
as you can see from the attached picture 1) how I can avoid cell aggregation. I have tried everything and still, it's frustrating because it affects my picture quality. What kind of neuron is N2a? Differentiation to which type?
2) for the protein expression, how can I locate it because it should bind so some ion channel
3) My labmate says that after the transfection under microscopy, they observe the protein expression moving( unlocalized). I did not face this. Is it possible?
4) i am planning to do a pH study to test how my protein is sensitive to pH change. I am planning to do this pH 6.6, 7, 7.2,7.3,7.4,7.5,7.6,22.214.171.124,8,8.5 what do you think? and how much should I wait after changing the pH for my cell to adjust to the new pH?
5) for preparing different solutions with different pH. Is it OK if I use my complete medium (which has FBS and antibiotics) and adjust its pH using NaOH and HCl? Is it OK or might it kill my cells?
6) as shown in the second picture, I think the GFP accumulates in the center. I believe is an uneven distribution. I may be wrong, so feel free to correct me. Also, after differentiation, I see that not only 1-2 cells that express GFP differentiated, and even if they do, it did not go to the axon. I was except that I will see some GFP on the axon because there are some ion channels.
i am starting working with insect cell protein expression from previously working with bacterial protein expression. for bacteria you can pellet and then freeze the cells after expression (and sometimes it even helps the lysis later).
I am gonna express a protein via Baculovirus in insect cells can i harvest and freeze the pellet before protein extraction and purification?
This study is fine. However, this finding will be
further strengthened If author confirm the protein expression of p-Erk, c-Myc,
Dicer-1 and BDNF by western blot.
N.B : I already did those parameters by ELISA and reviewers need confirmation by Western analysis
I am expressing a human RNA binding protein in E. coli and purifying it using ammonium sulfate precipitation followed by heparin, butyl, and size exclusion chromatography. While the first batch of protein purification did not show any RNA contamination, subsequent batches consistently exhibited RNA contamination. I have tried to maintain the same purification protocol but cannot seem to eliminate RNA contamination.
Have other researchers experienced this issue while purifying RNA-binding proteins, and if so, do you have any suggestions for troubleshooting or improving protein purity? I would appreciate any insights or recommendations to help me achieve a pure sample of my protein of interest.
Dear researchers, I am now trying to express several proteins from bacteria（Agrobacterium Tumefaceins) in one of its biosynthetic gene clusters. However, some of the proteins can be easily expressed and purified but many of them can not even be expressed. I also checked the pellet by SDS-PAGE and found that there were no over-expression bands in the pellet.
I want to ask how can I possibly get these proteins? Here are my protein expression conditions:
Culture until OD value reaches 0.6 and 0.5 mM IPTG was added(working concentration) to induce over expression.
Some of the proteins in the cluster can be expressed very well using this common protocol, yet others remain even unexpressed. I do not know what had happened, since there were no over expression bands in the pellet, suggesting over expression did not even take place.
I tried other recombinant tags like SUMO, MBP and GST, but none of them helped.
I am really stuck in this situation now. :(
If protein expression in blood monocytes was low (detected by western blot) and protein level in serum (detected by ELISA) was high? what dose it mean?
Note: the protein should not leave the nucleus because it’s a DNA-binding protein.
I have performed a bacterial protein expression, on a DNA construct, which I have sequenced and am sure is the target sequence. The construct should have been 42kDa however the post-induction sample shows an expressed protein band at 30kDa. What are the possible reasons for this? I would be grateful for your help.
I have tested a kit for cell free protein expression (Next generation cell free protein expression kit, wheat germ CFPS 700) from Merck and I didn't get the expected yield for protein production.
In the procedure of this kit you have to prepare DNA template by a game of several PCR, then in vitro transcription is realized from PCR template, and finally cell free translation using wheat germ extract.
All is good until transcription (agarose gel checking)
But after that the protocol is a mRNA purification using amonium acetate salt and ethanol.
I think these step is the problem because I loose a lot of mRNA.
Can somebody tell me if this step is necessary or if I can try to translate without mRNA purification? Or else, is there another methode for mRNA purification, that preserve its quality for the following transcription (the kit exclude phenol, trizol or ammonium sulfate purification that rendered mRNA unsuitable for translation)?
Asp was substituted for Glu. SDS-PAGE revealed that the yield of the mutant protein was lower than that of the normal protein and the yield of inclusion bodies in the mutant protein was higher than that of the normal protein. Bioactivity assay revealed that the mutant proteins were inactivated. I wonder if the amino acid substitution can induce a decrease in protein expression
1. Is TMEM192 the only selectable protein in LysoIP?
2. If I'm going to do LysoIP, should I make sure the TMEM192 protein expression do not change first?
3. After LysoIP, if I want to confirm other protein expression in the lysosomes, which protein can be used as an internal reference when LAMP1 increased after treatment?
We have been trying to insert peptide tags (<=100 bp) to a protein (~1,000 bp) in a 10 kb vector. The insertion was successful and verified by Sanger sequencing.
However, when the colony was liquid cultured, the plasmid concentrations were relatively low (100-200 ng/uL in 50 uL total for 10 mL cells compared to the original gene which is 200-400 ng/uL in 50 uL elution volume). And with a low A260/230 ratio (0.8-1.2 compared to the original gene ~1.4-1.8). A photo of the spectrum on nanodrop is attached. When we ran the plasmid on a gel (140 ng for all lanes), these constructs show around the same concentration.
Following, the same amount of plasmid (0.5 ug/500uL in a 24 wp, since the insert was small I did not bother to calculate the molar amounts) is transfected in mammalian cells (CHO/HEK) followed by 48 hour incubation. The protein concentration was analyzed by Western blot. Compared to the original plasmid, the protein concentration was very low (Western blot attached, left - original, right - after insertion). The same result was observed in immunofluorescence using two antibodies identifying the same protein. I know the plasmid is there and functional because one of the inserts was a nuclear-targeting signal and IF clearly showed localization to the nucleus (just fewer cells showing fluorescence). We're trying to analyze protein expression following tagging / mutations similar to site-directed mutagenesis, but I'm not sure this result is so believable in that it almost completely removed protein expression.
We also used the e coli glycerol stock on a new agar plate, and transformed new e coli (Dh5a cells) with the constructs with good growth overnight. But the plasmids were of a low quality (A260/230) and quantity (<200 ng/uL) regardless.
Are there any explanations for this? Or could I trust the western blot /IF result and say the protein was expressed less for other reasons, like degradation or a change of epitope? Thanks for any help.
if a protein has low intracellular expression levels, it is likely that the amount of protein secreted into the extracellular space will also be low?
if gene expression is low in specific diseases based on previous research,
But when I measured the protein expressed by the same gene in blood of same disease by ELISA I found high serum level?
what is the explanation for this difference between gene expression and its protein concentration in blood?
Differentiation of protein expression in mammary normal cells and mammary cancer cells and how to identify differences between normal and cancer cell lines with Immunohistochemistry.
I'd like to know that what are the different ways to know/identify whether a particular Gene is expressed or not ?
Few points from my side are :
1) identifying it's corresponding m-RNA transcripts level.
2) identifying the protein that was produced by the expression of that particular Gene.
Any other points ?
We have been struggling to get positive transformants when we used the commercial kit from TAKARA, Cat No: 3380, the included B. subtilis host strain is RIK1285. We have been following their protocol precisely which is available for online. We have further tried to manipulate the protocol by considering the recent improvements on B. subtilis expression, but still could not solve the transformation bottleneck. Did anyone already use this system or have any suggestion for the solution???
Thanks a lot for your answers,
I'm going to transiently transfect subcutaneous pre-adipocytes with His-tagged plasmid DNA [pcDNA3.1(+)] containing ADIPOQ gene with a SNP to study the adiponectin expression. Can any expert shares to me what is the best lipid based transfection reagent to transfect pre-adipocytes; FuGENE 4K or Lipofectamine 3000? How long is the most optimum waiting time after the transfection process to proceed with protein expression? I'm also going to treat the cells with few drugs, and will perform RNA quantification (qPCR) and measure the level of protein (adiponectin) via ELISA. Should I perform western blot as an analysis to confirm protein expression before I proceed with drugs treatment after the transfection done? Million thanks in advance for the ideas and generous support.
I wonder why E.coli cell pellet freezing is recommended before the lysis during protein expression and purification process.
Most of the commercial protein purification protocols recommend it,
and I also have some experience that fluorescent protein-expressing E.coli pellet showed more vivid color when it was in the freezer longer time. (though I didn't quantitatively measure it)
I've tried to find the regarding report but I couldn't.
Does anyone knows the reason or have idea?
Thank you for your opinion in advance.
Does anyone have experience with "missing terminal nucleotides" after sequencing? I use NEB Q5 polymerase, as I know it has strong 3'- 5' exonuclease activity.
So far I had no problem, but now I have two cases
1. Site directed mutagenesis on a 10 kb plasmid. It's nicely mutating the desired site, but a part corresponding to the 5' end of For primer is missing after sequencing (After DpnI digestion, ligation, transformation). In this case I used non-overlapping primers as proposed (currently i'm trying with overlapping primers also)
2. It's a 2.5 kb plasmid, so far I did similar nucleotide substitutions with succes. The problem is the same: a nucleotide normally present at the 5' end of the For primer is missing when i'm sequencing. oK? It's a "T" so maybe it was just not the best choice.
Do you have any idea how could I avoid these missing nucleotides?
Dear all colleagues,
I am new in the cell biology field. Could someone recommend me the technique or method to investigate the mechanism between pathogen and protein expression (of the host)?
Thanks in advance
Hi, I produced a stable expressing Cas9 cell line. After transducing the cells, I checked the protein expression of my protein of interest and I understood that the transduced cells expressed a protein of interest less that the non-transduced cells. Does anybody have the same experience?
Does anyone in the community have experience cultivating Expi293F™ Cells Thermo, A14528 under SILAC conditions? They are usually cultivated with their special medium to achieve high protein expression rates. However, there are only the commercially avaible standard media DMEM, RPMI, etc... for SILAC. Any ideas or experiences?
I want to compare the protein expression level of different bacterial transcriptional regulators in E. coli. Are the GFP or other fluorescent protein genes suitable for determining protein expression? I have tried western blot, but the protein bands are too weak when using His tag. any suggestions?
Hi, I need to create a bacterial strain with 3 vectors, it already has 2 and I need to add one more, but I don't know how to choose (2 vectors are for protein expression). I know they cannot have the same ORI site, but when I go through the addgene database it seems to me like all have the ori from pBR322. Can you give some advice regarding your own experience with expression systems construction or do you use any other database for plasmids? Thank you.
I have been studying DLL4 protein expression on HUVECs. But, every time my membrane shows many bands. I have been using a polyclonal antibody at appropriate dilutions. Is there any specific antibody that works for others? Thanks in advance
The pET-17b vetctor (the enclosed file) lacks the lac repressor expressing gene, and also seems to lack a lac operator in the upstream of the insert site. However, a lot of literature used pET-17b to express protein with IPTG induction. I am wondering how pET17b could be induced or enhanced for protein expression in this way.
I am trying to express and purify nanobodies from a nanobody library in E. coli. I am using a pBAD vector to which I added the pelB sequence for periplasmic localization. The nanobody is fused to YFP with a His tag.
I induce the expression with arabinose and I can see on an SDS-PAGE gel that the nanobody is being expressed (although the expression seems to be lower than for some of the other proteins that I am expressing using the same vector), but I am having trouble with the extraction and purification steps.
I have tried to extract the nanobodies using lysozyme with PMSF following a protocol that usually works for me and I have also tried the osmotic shock protocol (
I do not have any experience with nanobodies so maybe there is an important step in the protocol that I am missing or not doing properly. I would appreciate tips or good protocols for expressing and extracting nanobodies.
Hello, I have a problem with the production of recombined, quite small (138 aa), protein in T-REx HEK cells. Plasmid was sequenced so that's not the issue. I have used this plasmid with other proteins therefore its functional. Protein contains signal peptide (checked in other contsructs - working well) and HisTag at N- and C-term ends. I've tried to find the produced protein in medium and cell pellet using gel staining and WB with anti-HisTag ab as well but with no result. The protein is composed of glycine in ca 40%, so I've started wondering wheter it might be a reason of such miserable effect?
I'm writing because protein expression is not working well.
I'm using is a promoter called HSP60 or CJ1 in C.glutamicum, but it doesn't express well, so adjusting several variables (ori, backbone, terminator, etc.), but I think all conditions are fine. So I think there's a problem in the process of expressing it.
What I'm curious about now is that this promoter is Constitutive expressed, so there is no special process, but I'm posting this because I thought it might be a promoter that needs to shift on the temperature.
Is there anyone like me who used this promoter to express it?
I'd appreciate it if you could leave any comments.
Suppose you are trying to express a protein in Bacteria but that protein is coming in Inclusion bodies but not secreted out. So is it possible if I express that same protein in mammalian cell with signal peptide to make it secreted out in the supernent so that I can purify it easily ?
I've few queries regarding bacterial and mammalian plasmids for expression of Gene of Interest. What plasmid elements/components that are differ between bacterial and mammalian Plasmids to express a gene of Interest.
According to me :
The elements/components that are common between bacterial and mammalian Plasmids are :
- Bacterial ori of replication.
- Bacterial selection marker.
- Promotor + gene of Interest for Expression of Gene.
The elements/components that are differ between bacterial and mammalian Plasmids are:
- Mammalian Ori such as EBV or SV40 if the Transfected cells expressing the Epstein–Barr virus (EBV) nuclear antigen 1 (EBNA1) or the SV40 large-T antigen for Episomal replication of Transfected plasmid.
- Mammalian selection marker (For positive selection of cells that take up plasmid).
- Promotor + gene of Interest for Expression of Gene + PolyA (example SV40 pA or CMV pA)
- Reporter Gene.
I'd like to know is there any other differences?
I'd like to know the Maximum Yield we can achieve with CHO cell line (Irrespective of CHO-s/ExpiCHO/CHO-K1 e.t.c and also the mode of operation like batch, Fed-batch and Perfusion) ?
Hello! I'd like to ask a question about protein expression. My question is how I can remove N-terminal formil-methionine from E.coli recombinant protein if the next amino acid after methionie is phenylalanine. It is known that methionine aminopeptidases such as MAPs from Pyrococcus furiosus are depent on a penultimate residue of substrate and they do not react with Cys, Asp, Asn, Leu, Ile, Gln, Glu, His, Met, Phe, Lys, Tyr, Trp, Arg amino acids. So, How i can deal with this problem. What type of enzyme will be applicable for M↓F removal ?
I have a proteomics data and want to compare male-female mice protein expression levels. I need to find differentially expressed proteins between two group. Student's T test just compares the mean values and gives a p-number. But I want to find which protein's expression level is significantly upregulated or downregulated. So what test/method should I use?
Thanks for any contribution.
I will be synthesising single-stranded linear and circular mRNA by an in vitro transcription reaction (T7 HiScribe IVT synthesis Kit, NEB) followed by DNAse 1 treatment and mRNA purification. This mRNA will be lipofected into mammalian cells for protein expression. What would be the best way to preserve this mRNA for mammalian cell work? What would be the optimal vehicle for freezing? (Nuclease-free water/TE buffer/DPBS?)
At what temperature should mRNA be stored? Would liquid N2 work or -80C will be sufficient?
How thawing would affect this mRNA? How much of single stranded mRNA will be degraded upon thawing and is there a thawing routine for preserving single-stranded mRNA?
I would like to know whether anybody ever found discrepancy between RNA and protein expression (for the same cell line in culture), by PCR and immunocytochemistry, respectively.
Specifically, while the immuno seems to identify the protein which is also, apparently, highly expressed (the antibody has been doubled-checked on a cell line used as positive control and known to express the protein), its RNA is barely detectable by PCR amplification (I would say is not there).
Logically speaking, it's impossible to have protein without RNA. I thought the immuno is possibly an artefact, but the same antibody identifies specifically the protein on control cell lines.
Can someone please give me suggestions? Thank you
I'm using E. coli BL21 (DE3) GroEL/ES cell for my protein expression by IPTG at 0.1 mM concentration at 18 degree Celsius temperature.
I'd like to know that the Signal peptide should be synthesized in which terminal (C- or N-terminal) of a protein in order to secrete out extracellularly and why ?
Does anyone know if the high-copy number plasmid is available for protein expression? I modified one plasmid from a low-copy number to a high-copy number by exchanging the ori, I do not know if this change could influence the protein expression.
In my project I use the pET21a(+) expression vector to express a bicistronic gene. For some technical issues we got in the past the construct without T7 promoter. Today, I have the correct system with the promoter. I decided that it could be interesting to compare the activity of my system to the one without the promoter as a negative control. Surprisingly, I got a parcial activity of the "no promoter" control that could be explained by a basal expression of the proteins. I wondered if this could make sense.
I want to use tetracycline as selection marker for my next experiments. I was advised to perform the experiments in dark. I want to know if this precaution is really required. I haven't found any literature supporting light-sensitivity of tetracycline. Please help...
Thanks in advance... :)