Saccharomyces cerevisiae - Science topic
Saccharomyces cerevisiae is a species of the genus SACCHAROMYCES, family Saccharomycetaceae, order Saccharomycetales, known as "baker's" or "brewer's" yeast. The dried form is used as a dietary supplement.
Questions related to Saccharomyces cerevisiae
Looking for a straightforward approach to isolate nuclei that does not require the use of an ultracentrifuge... we generally work with mitochondria, however, we are interested in examining some nuclear proteins. Our crude nuclear pellets from our mitochondrial isolations appear to be contaminated with broken mitochondria. Any suggestions would be greatly appreciated!
I am trying to knock out a gene in 2 strains of S.cerevisiae, BY4743 and W303 with CRISPR/Cas9. I am using the same gRNA and donor DNA for both strains because I want to knock out the same gene the same way. There is no alteration in the sequence which should be complementary to the gRNA, I checked. It works with W303 nicely, but not at all with BY4743. What can be the reason?
(Both strains are haploid.)
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'm working with a Saccharomyces cerevisiae strain that tends to form big cell aggregates. In order to take this strain to a flow cytometer, I need to get single cells, but although I've used up to 250 mM of EDTA and strong mechanical forces, I've been unable to disaggregate the yeast's chains. Does anyone know a way to make cells separate?
I have a fluorescent protein (excitation - 497 nm) expressed on the surface of yeast, however as the cells are auto fluorescent, I am unable to see the expressed and non-expressed populations in flow cytometer as well as microscopy.
Note that I cannot use a dye in the red area (excitation - 640 -660 nm) as I am using this as a tag to measure another expression and I have no issues with this antibody as I could clearly see two different populations
Any inputs on reducing autofluorescence in saccharomyces cerevisiae would be very helpful. Thanks in advance!
I’m trying to reproduce the library transformation efficiencies seen in this 2010 paper:
in which they claim 1.5 x 10^8 cf/ug DNA. My intact circular vector is the same size as their pcr product + linearized vector, but I’m getting 10^5 ish transformants/ug, similar to chemical transformation with the zymo kit.
Has anyone successfully done this? so far we’ve tried different electroporators, voltages, recovery media, etc but nothing seems to be working.
Any other ideas or troubleshooting would be much appreciated, thanks in advance
I'd like to know what other method can be used to assemble 3 different DNA sequences, apart from the recombination method in saccharomyces cerevisiae, Gibson assembly, and PCR assembly?
I will ferment ulva substrate with saccharomyces cerevisiae yeast. Can I add it directly or do I have to do the activation process first? If there is a need for the activation process, can you help me with the procedure?
When purifying total RNA from yeast that have grown on nitrogen-limited medium, it always appears more degraded when coming from stationary phase than from log phase.
RNA coming from log phase always looks fine.
I'm using a modified protocol from A rapid and simple method for preparation of RNA from Saccharomyces cerevisiae (Schmitt et al., 1990).
Thanks a lot!
Combining feed enzymes, and minerals with probiotic microbes will affect the efficiency of microbes??
How to make the Composition by mixing the 3 of them?. (Probiotic, Feed enzymes, Minerals)
I am trying to design guide RNAs for cutting the yeast 2 micron plasmid using CRISPR. Using Chopchop, I got a 23 nt sequence which was suitable for gRNA design.
When I blast the 23 nt sequence against the S. cerevisiae genome to check for any off-site cutting, I get multiple results of 100% query coverage, see the attached picture. However, when I click these results to investigate the alignment further, the alignments are all 14 nt or shorter. It seems that BLAST is finding multiple short alignments which cumulatively cover the entire query sequence - however, none of the individual alignments match the entire query sequence, which is what I'm interested in.
Why do I not just get the highest query cover result as being 14/23 nt=60.86%? And is there any option I can toggle to get these values?
We've been extracting RNA from S. cerevisiae samples grown in synthetic media and in our RNA screentape data we see a large peak below 200 nt that keeps appearing. The RIN values and purity of the samples are all fine. The samples also pass the QC for illumina sequencing and subsequent library preparation. RNA was isolated using an acid phenol/chlorophorm/IAA protocol. At first i thought this was degradation, but it the rRNA peaks seem fine and it doesn't look like your typical degradation profile. Does anyone know what this peak represents?
Hello guys, I am going to start cloning experiments with Saccharomyces cerevisiae. I need a standard protocol for isolating Genomic DNA from Saccharomyces cerevisiae. If anyone of you are working on yeast, kindly help me out.
In my project I am measuring the survival rate of the yeast cells after in-vitro exposure to gastrointestinal conditions.
Previously a masters student used Plate Counting to determine the survivability of the strains and observed 4 % of the survivability.
Later I run the same protocol in another date and used Flow Cytometer to determine the survivability. I did not use a stain for the live cells, but only used propidium iodide for the dead cells. Positive and negative controls showed the clear difference between the cells from the fresh culture vs cells exposed to 70% isopropanol for 1h.
In my experiment, 99% of the cells were in the "alive" zone.
I am trying to find resources to understand why there is such a big difference in these two methods. Could it be VBNC, something regarding the stains or something else?
Dear yeast biologist,
I performed 4-5 time yeast transformation in S. cerevisiae SK1 MATalpha strain but I am unable to get transformants.
I performed and checked all the necessary steps as per my knowledge.
If possible please suggest me, how to troubleshoot the problem?
I will transform the plasmid that contains the gene of interest. The plasmid will be transformed into yeast cells by electrophoration. The yeast used comes from isolated baker's yeast to obtain a single colony. is it possible for the plasmid to enter into yeast cells? or do I need a competent yeast with a specific strain specifically used for transformation? Please help me.
I have been trying to stain my yeast cells with DAPI after fixing them with 4% PFA and I am not getting a good result. Maybe someone has a tip they can share on how to do it to get nice pictures!
Also, is there a way to use DAPI staining in cells in stationary phase without having to add additional glucose to the media??
Thanks a lot for your help!
Please suggest me why I am not able to get transformants and how can I troubleshoot the problem?
Initially when I transformed and selected on Hygromycin and G418 plate, I got 5 to 15 colonies per plate but now not a single colony I observed after 5 rounds of transformation.
I checked the incubation and heat shock temp.
I prepared the transformation reagent freshly, such as, TE-Lithium Acetate buffer and PEG.
I used DMSO to increase the transformation efficiency.
I maintained proper cell density during the log phase culture.
now what I should try????
I am planning to do the transformation of Yeast integrated plasmids into a lab strain of Saccharomyces cerevisiae. Till now I did transformations of plasmids like pET43a and pET28 into DH5alpha cells and BL21 cells of E.coli. If anyone of you did the transformation of plasmids into Saccharomyces cerevisiae, could you please share a standard protocol?
Howdy, I have been trying to express a big protein (~1500 AAs) of bacterial origin in baker's yeast which needs to get into the nucleus. The protein is fused to E2-crimson fluorescence protein, and SV40 NLS is used. But fluorescence microscopy revealed that only a small fraction (~10%) of yeast cells had red signals in nucleus. The picture is attached.
I wonder why nuclear localization is so inefficient, and any suggestions to improve are welcome!!
We performed transformation on Saccharomyces cerevisiae, where we introduced a gene mutant library (roughly 14 mil. mutants). Because of the high transformation efficiency, many colonies appeared and created a lawn (the attached picture displays 1 out of 20 transformation plates).
It is very important to be able to calculate the exact transformation efficiency in order to estimate the size of the library.
Does anyone know a way to count the cells despite the 'lawn' formation?
Also, does anyone have any suggestions on how to stock the library?
(could it be as simple as scraping all the cells of the plate and making a glycerol stock?)
Thanks a lot for your help!
I'm currently confused on the centrifuge speed needed for separate yeast cells (Saccharomyces cerevisiae) from its aqueous medium
I need the yeast cells to still alive to ferment herbal extracts
I am curious whether I can culture mammalian cells and yeast cells in the same culture room. Our lab has one cell culture room (walls are made of something similar to glass, not actual walls) that is the shape of a rectangle. Inside this room, there is another small room with its own door.
We would like to culture yeast and mammalian cells, but we are scared of contamination of the yeast onto the mammalian cells. For the yeast culture, I wouldn't be using a biosafety cabinet, and they would be cultured in a separate incubator.
I was planning to use the small room or yeast cell culture, and the bigger one for mammalian cell culture. The culture tools wouldn't be shared in any case.
We are the most worried about airborne contamination. For example, we are scared that if I handle yeasts first in the small room and then change my gloves to handle mammalian cells, some yeast cells on my lab coat or my skin would contaminate the mammalian cells. Or also when the door between the two rooms is open when workers come in or out.
I would like to know if this kind of contamination is possible, and if, with aseptic practice and maintaining tools isolated we could still culture them in the same "room". Also, may an HEPA filter/air purifier solve the problem? Any suggestions and solutions are more than welcome!
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?
I'm doing a yeast display experiment and have been sorting yeast cells (S. cerevisiae) with a BioRad cell sorter.
In the later rounds I've started getting contaminants on the plates. Some plates do not have any contaminants at all. In other instances it's quite possible that the contaminant is being passaged along with the sorted cells (although I'm gating a double positive population...)
The colonies have a raised margin and in some instances a "button" in the middle. The attached pics have a 200 ul pipette tip for scale; there are also some typical S. cerevisiae colonies visible in the images.
Plates: synthetic complete - trp, dextrose, pen-strep. ~2 weeks old and stored at 4C.
Growth: 2 days at 30C. Incubator is also used for E. coli plates.
Yeast strain: S. cerevisiae BJ5465
Plasmid: gal promoter so there should be no cell surface display of the DARPin library.
Cell sorter: BioRad S3. The sorting chamber is not sterile. Only yeast cells have been sorted recently although some Corynebacterium cells were analyzed in the same time frame (different days though) that yeast were sorted.
Thanks for any ideas!
Hello, I'm trying to make media for S. cerevisiae that contains everything it needs to survive except any kind of sugar. The goal is to make stock without sugar then add in specific sugars of our choice. This is to research a gene believed to function in the metabolism of complex sugars, so we need to starve the yeast of all but specific complex sugars to test if that's actually its function.
Does anyone know how to essentially make YPD from scratch so that we have the option of leaving out any sugar containing ingredients? Thank you!
Im working on a project and need to make sure this rough draft of a protocol is valid. Is this a good outline, am I missing any steps?
Im basing this protocol off this paper:
A Cas9-based toolkit to program gene expression in Saccharomyces cerevisiae. Reider Apel A, d'Espaux L, Wehrs M, Sachs D, Li RA, Tong GJ, Garber M, Nnadi O, Zhuang W, Hillson NJ, Keasling JD, Mukhopadhyay A. Nucleic Acids Research. 2016 November 28; DOI: 10.1093/nar/gkw1023. PubMed PMID: 27899650.
- Acquire dna and crispr plasmid through 3rd party supplier
- purify dna
- amplify dna
- co transform yeast with dna, cas9 crispr plasmid and transformation mix
- incubate yeast
- examine yeast to confirm transformation
I use S. cerevisiae for yeast display and I suspect I have a contamination in my cultures but it is not obvious and I don't currently have a microscope in the lab.
I grow in minimal media selecting for Trp auxotrophy complementation in the transformed yeast.
Recently my yeast cultures don't seem to display and they are growing slowly and have a less pungent smell compared to S. cerevisiae. They don't smell like E. coli either and colonies on plates have same colour and shape as S. cerevisiae as far as I can tell.
What are the common contaminants yeast scientists see and how to deal with them?
I know about Candida... is microscope the only way to identify a contamination from Candida or other species?
I would like to know the culture collections or other private institutions who could provide yeast strains for protein production possessing strong promoters, selection markers and possessing other properties of a strong expression system.
I built a fusion protein with GFP and am trying to confirm it via fluorescence microscopy and WB.I can see the GFP signal (nuclear localization) in roughly 80% of my budding yeast cells.
However, trying to confirm by WB and am getting no bands using anti GFP. I have increased my primary antibody concentration and included a positive and negative control as well. The loading control also shows a visible band for my samples. The size of protein is around 160 kDa and my positive control is 120 kDa, so I don't think transferring to the membrane is an issue.
If it matters, I use a standard alkaline lysis method for protein extraction.
I am trying to ferment the enzymatic hydrolysate I obtained after pretreatment to produce bioethanol. So, as a trial, I did the experiment in a fermenter using YPD and checked ethanol production using Saccharomyces strain purchased from a Supermarket locally. The yield I got was very low and the glucose consumption rate was also low. Can somebody please suggest where I can buy high-efficiency yeast strains from India that can be used for fermentation?. It would be a great help!.
Thanks in advance!
I'm using the QIAGEN Multiplex PCR Kit and 10 primer pairs (product sizes from ~130 to ~900 bps) to monitor mitochondrial DNA in Candida glabrata, a close relative of Saccharomyces cerevisiae. One primer pair target the genomic DNA, and is a positive control in case the mitochondrial DNA is completely lost (this is possible in yeast). The other nine amplify nine different genes on the mitochondrial DNA. Every product has a distinct and fixed size, and they separate well in a 2% agarose gel.
Every primer pair works fine when run alone, but when run together (for 45 cycles), I only get 7 bands. The 3 missing bands are the genomic DNA product and two of the mitochondrial rRNA subunits (SSU and LSU). The 7 bands that do work are all protein-coding mitochondrial genes, which suggests a pattern, although I don't have an explanation for it.
A number of observations that might be relevant:
1) In C. glabrata, the mtDNA is present in a much larger copy-number than the gDNA - I estimate about 20-50 times more, but I could be wrong. Still, given that PCR is exponential and I'm using 45 cycles, I should still get a band from the gDNA primer pair, even if weak.
2) The LSU and SSU products are quite GC-poor (20-23%), but the other mtDNA products are not much better (~24-32%), so I don't know if that matters.
3) All the primers were designed to have a similar Tm.
4) Running the PCR in a temperature gradient didn't seem to do much.
5) In rho- petites (yeast that completely lost their DNA) I see no bands from the mtDNA primers (as expected) but the gDNA band does appear (also as expected - perhaps because the mtDNA primers don't interfere with the reaction anymore?).
I can start playing around with the primers and their mixes, but I feel there's something theoretical I'm missing. In any case, any and all advice will be appreciated.
Thank you in advance!
Corn stover is a cheaper lignocellulose that is used to produce biofuels. There are numerous methods for pretreatment of corn stover before hydrolysis. Among those, which one is the most efficient?
I completed a fed-batch fermentation with the addition of spent sulfite liquor (SSL) (a waste stream produced in the paper and pulp process) to S. cerevisiae that terminated with a 60 (v/v) % SSL concentration. Prior to this I completed a batch fermentation with 60 (v/v)% SSL, where I incubated the yeast (in YPX) for 24 hours prior to the addition of the SSL. The yeast was incubated in batch mode (in YPX) for 96 hours before the addition of smaller volumes of SSL (for the fed-batch fermentation).
The ethanol concentration only increased by 3.10 % from the batch mode to the fed-batch mode. I was expecting a much larger increase since the fed-batch mode conditions the yeast to the inhibitory compounds present in SSL. A previous study conducted managed to obtain a 50 % increase in ethanol concentrations using the same substrate. The difference is that the initial batch mode for the fed-batch fermentation was only for 48 hours as opposed to 96 hours (as in my study).
Is there any explanation for the drastic difference in the results obtained and the poor performance in my study?
Hi, I am wondering if there is a correlation between colony size on a YPD plate and fitness. I am working with S. cerevisiae.
Also, does the fitness calculated with the size of the colony also correlate with the fitness calculated from the growth on liquid media?
#yeast #saccharomycescerevisiae #fitness
does someone know a paper where authors engineered the arabinose induction system of prokaryotic organisms in yeast Saccharomyces cerevisiae, like an IPTG induction system, for orthogonal gene expression?
I am working on the disruption of some yeast genes to create new strains... for my research project "Metabolic Engineering of Saccharomyces cerevisiae as lipid cell factories for Bio-Diesel production"
I work primarily with S. cerevisiae and commonly perform western blotting on total protein lysates. Our lab uses a fairly standard TCA/Bead lysis method (https://www.med.unc.edu/pharm/dohlmanlab/resources/lab-methods/tca/) for extraction of total protein from cells. I base the re-suspension volume after lysis on OD600 (10 µL SDS Loading Buffer/10^7 cells), however, I am having difficulties getting optimal loading using this method (sometimes the loading is fine, whereas other times there is a large variation between samples of similar OD600). Does anyone else use this method and have tips for troubleshooting or can recommend an alternative method?
There are several known long non-coding RNAs (lncRNAs) like H19, Xist, NORAD and Malat1 in mammalian cells. What, your opinion, are in the most important examples of functional lncRNAs in Saccharomyces cerevisiae that exert their function via the RNA molecule not via the act of transcription happening at their genomic locus?
I was wondering if anyone here has some insight into what should be the distance between the promoter and the translation start site / initiator-ATG when making a DNA construct? In particular, I`m interested in this in making an S. cerevisiae expression plasmid. And I`m for example wondering how much of a multi-cloning-site I could put between the promoter and ATG.
One of the critical issues seems to be how "promoter" is defined. Should I look for specific sequences here (e.g. TATA box; does anyone know the sequence or a regular expression to look for them?)? The same issue for the translation start: Do I just look for the first ATG?
Hi everyone, I am hoping to take a look at the cell cycle progression for my budding yeast and I have chosen alpha factor as the method of cell cycle synchronization. I wish to ask for your help because one of the mutant I am currently working with simply won't arrest in response to alpha factor.
The issue: mutant unresponsive to alpha factor, after 150mins arrest, only 50% of the total population are shmoos (~80% unbudded); when I took a look at the cell culture with flow cytometry (PI stained), a lot of cells seems to be stuck between G1 and G2. When the same alpha factor concentration is applied to WT, it arrest as expected.
Experimental condition: I used buffered YPD media (pH~5). One the day of the experiment, I diluted my cell culture (~1.5 OD600) back down to 0.1OD, allow it to double once before adding the alpha factor for arrest.
Optimization I have attempted: I have played around with the alpha factor concentration and arrest time but strangely none of them seems to affect the arrest efficiency. I still get one peak between G1 and G2 for my mutants.
Therefore, I wish to check in with everyone on research gate to see if someone can give me some pointers for how to further increase my alpha factor arrest efficiency.
Please let me know if there's any clarification or additional information I can provide. Thank you in advance!
We have this older plasmid YEpTOP2GAL1 for expression in S. cerevisiae. We don't have the plasmid sequence which would be necessary to design primers or new cloning constructs. Does any one have the sequence for YEpTOP2GAL1? I can't find it in any references or sites like Addgene.
References - PMID: 24124932, PMID: 23129636
I have to transform two plasmids into yeast. One is 8kb with trp marker and other is 14 kb with ura marker. both are 2micron. I used Gietz et al method.
First, i tired tansforming both plasmids at same time and after plating on trp/ura plates, no colonies were observed.
Secondly, I prepared competent cell containing 8kb plasmid (trp marker) and tired transforming 14 kb plasmid (used 500ng). Still no colonies were observed. But the controls were perfectly alright. i could see lawn of cells in Trp drop out plate (POSITIVE CONTROL). As negative control, cells were plated in ura drop out plate and no colonies were observed. How could I solve this issue?
I am working on Phosphoproteomics experiment by using SILAC (on S. cerevisiae). I have got back my MS data and was suggested to use Proteome Discoverer 2.2 to analyze the data. Since this is my first time doing Phosphoproteomics, I need to get some advice (and direction) to analyze the data. Does anyone know what is the first thing to look for? (To my understanding, I need to look up the Heavy/Light ratio for the heavy-labelled and light-labelled peptides in order to quantify the protein abundance)
Any inputs are appreciated.
I work on Budding Yeast (S. cerevisiae), and often face hurdles and issues with techniques, protocols and sometimes I am just plain curious. I was wondering if there is any exclusive platform for yeast biologists to come together and discuss their research related problems and queries and help each other. I am sure that we learn so many things from our mistakes, and sometimes the smallest thing is actually what is needed for an experiment to work. Can you guys please suggest or share a link if you know, or are a part of such a group. I would love to be a part of the same.
What are the names of strains of Saccharomyces Cerevisiae which are generally used in production of nutritional yeast?
I have been using Takara's Yeast Transformation System 2. Using the kit's protocol, I transformed Saccharomyces cerevisiae with plasmids that have auxotrophic selection (trp and leu) in them. I did spread 100 ul of 1/10 and 1/100 dilutions of transformed cells on respective SD/-trp and SD/-leu plates. After 5 days of incubation at 30C, nothing appeared on the plates.
I was wondering, if after transformation and recovery in the YPD Plus medium (provided with the kit), if I wash the cells with 0.9% NaCl and grow for 5-8 hours in the SD/-trp and SD/-leu broth, before plating on the SD/-trp and SD/-leu agar, will it make a difference?
Kindly also let me know, what all of you look to change when your transformations fail.
I am searching information about increasing yield of secondary metabolites that are heterogeneously produce in S. cerevisiae.
First I transfer several genes into yeast, then I wanna to optimized the yield of these metabolites. I wonder that whether epigenetic modifications play any roles in enhancing the production of these metabolites in the yeast transformants?
Thank you very much!
I am trying to precipitate an endoglucanase from liquid culture of saccharomyces cerevisiae. the protein is under control of Gal1 promoter and I already confirmed that the endoglucanase is expressed by using CMC agar plate and confirmed formation of halo.
I read several references about precipitation with acetone or TCA or ammoniums sulfate and I would like to chose one for precipitating my protein and maintain its functionality because I will use it in enzyme assay.
it is not possible to precipitate the protein through use of column because of limited funds.
thanks for your help.
I performed qPCR analysis using the FungiQuant primers to target the 18S region for fungi to determine fungal biomass. I used a plasmid standard containing this region from S. cerevisiae and was able to get some biomass measurements. However, I am unsure how to account for variation in gene copy numbers. If anyone has any ideas or can guide me to an article, I would greatly appreciate it.
I am maintaining yeast cultures in bulk. Since, yeast's life cycle is significantly slower than bacterial. I was wondering if I could use some antibiotic which can prohibit the growth of bacteria without interfering the yeast's growth.
Any suggestions will be welcomed.
I am trying to confirm the enzyme activity of a cloned endoglucanase gene under inducible promoter gal1 which is transformed into the S. cerevisiae. first I confirmed the expression of the endoglucanase through use of CMC plate and red-Congo staining in comparison to control (empty vector).
my question is when i will do then induction to express the gene, I should use galactose and the DNS assay quantify the free sugars, so the supernatnt will contains galactose and the DNS-assay will not allow correct quantification of free sugar?
I'm researching the growth law of budding yeast Saccharomyces cerevisiae. I'm searching for growth rates under different conditions like different culture media(YPD, SC, etc) or different temperature, but I find it diffcult to obtain these data. Most papers just mention that they harvest log-phase yeasts when OD600=0.5 rather than provide the growth rates. Are there any databases or papers providing these informations? I think these information may have been systematically reported long time ago because this question seems very fundamental, so please forgive me if this question is too naive.
I am trying to express a cellulase gene from an other yeast in saccharomyces cerevisiae YPH252 and i cloned it into the pyes2 vector. transformation was done using electroporation and selection was done using uracil- and synthetic media (SCU). after that I transferred the colonies to SCU media and grew the colonies for 2-3 days.
I tried the colony pcr for yeast using SDS, NaOH, distilled water, incubate at 99 C for 10 min, vortex and spin down.
I used 1-2 ul, however the pcr was negative without any bands except the positive control (plasmid as template).
second part of question, is it possible to use the strain YPH252 for galactose induction?
I'm working with yeast strains with a S288C background that display a low-efficiency sporulation profile. Thus, it is needed to select spores after sporulation and restrict the non-sporulated cells. The strains I work with do not allow for carrying this step out in a genotype-dependent way using selection markers. I have come across some protocols that suggest using enzymatic activities such as zymolyase, glusulase, as well as a protocol suggesting the use of diethyl ether. Are there any other methods/protocols you are aware of? Would you suggest any of the above?
Resently, the result of acetic acid boosts the yield of ethanol using Saccharomyces cerevisiae fermentation under the single carbon source of glucose was obtained, which was different from most of the report. can anyone give me some advice?thanks.
I want to do a plasmid shuffling assay in saccharomyces cerevisiae YPH499， Which series of plasmids are suitable for YPH499, 3 or 4? Is there any difference between series 3 and 4? thanks very much
I'm trying to separate dividing and nondividing yeast cells from my medium using flow cytometry. I know that nondividing (G0) S. cerevisiae cells are between 4.1-4.9 um in diameter, but I can't find any resources showing the diameter of these yeast during G1. If I knew about what diameter they should be at the end of G1, it would be much easier for me to predict the kind of markers they'd be showing through flow cytometric analysis (using forward light scatter, which is correlated with cells size).
If you've ever worked with these cells, do you have any approximation of their diameter at the end of G1?
I have a problem with protein expression and the purification of genes from a plant source in yeast. I cloned the gene successfully in pYES2.1 through TOPO TA cloning kit (Invitrogen) and after sequencing, I have transformed recombinant plasmid in INVSc (Saccharomyces cerevisiae). I tried protein expression for this candidate by galactose induction (2% working conc.), but the protein is not getting expressed as I am not getting any specific band on the SDS page at the desired location. All I observed were bands of internal proteins (yeast). I tried to amplify the candidate gene from transformed yeast cDNA..and it was there in the induced yeast as compared to the non-induced one. Does anyone have any suggestions..why the protein is not being expressed and is there any purification kit recommended for the same?
Kind regards, Alka Jangra
Why it is a common practice to use S. cerevisiae alpha-glucosidase instead of the human enzyme, for in-vitro and in-silico/docking enzyme inhibition studies? Even the sequence of the yeast and human protein is not exactly the same.
I needed the molecular weight data for Saccharomyces cerevisiae, but I couldn't find the data and the molecular formula. Please help me. Thanks.
I read somewhere that the highest it could reach 10% but I could not be sure as I could not find the source anymore. Thanks!
#bioethanol #fermentation #saccharomyces cerevisiae
According to the MycoBank, the taxon "Saccharomyces cerevisiae var. bayanus" and "Saccharomyces cerevisiae f. bayanus" are not valid. But several commercial products are composed by fermenting yeasts classified in this way. If these are not valid taxa, which one is correct for the strains in the commercial products?
Thanks in advance.
I am analysing RNA-Seq data and wish to know what is the chance of a high quality mapping of a read (with high quality base calling) to two different ORFs. Therefore, I am trying to find the genes with highest degree of homology within S. cerevisiae genome. So far, google and pubmed have not been very helpful. I know they analysed this in this paper -
I cultured Saccharomyces cerevisiae in YPD broth and incubated different flasks (some of them treated with a vegetable peel)in a shaker water bath for 48 hours at 30c. the untreated yeast cultures were ended up with a white layer of yeast growth at the top of the flask. it was tough enough that I couldn't homogenize the samples. I want to measure yeast growth indifferently treated flasks. Is there any way for preventing such a layer? or measuring the yeast growth correctly?
I have tried to use TTC agar overlay method for a few times to identify respiratory-deficient yeast which were affected by EthBr. The problem is that cells which should have stayed white also have turned red or pink.
Evaluation of the overlayed colonies was done after 1 h, 2 h, 3h and 6 h. I tried different TTC concentrations: 0,1%, 0,05%, 0,03% and 0,01%.
I have attached photos of the colonies which were affected by EthBr and which were not.
We are conducting a measurement of Ribonucleic acid from baker's yeast (S. cerevisiae) (Sigma-Aldrich R6750) concentration using Eon Biotek Microplate reader. We dilute the RNA in various concentration (ranging from 100 microgram/mL until 1000 microgram/mL) using deionized water. Then we move the samples to a 96 UV-Vis plate and calculate the absorbance of the samples using Eon Biotek Microplate reader in lamda 260 nm and 280 nm. The results are quite confusing, first get the trend by the increase of concentration, the absorbance also increase, but then when we tried to calculate the RNA concentration using:
[concentration] = 40 x Absorbance 260 nm x dilution factor
we cannot get the right result,. Then when we calculate the A260/A280 ratio, the values are below 1, which can be an indication that there is too much protein in our samples.
My question is where the problem come from? Is it from our procedure, the purity of the RNA, or the setting of our microplate reader?
We are doing western blot with yeast protein extracts. We are applying a wet transfer method using Tris-Gly as a transfer buffer (with 10% methanol) and running it with constant current (400 mA) for 2 hours at 4°C, while the tank is put in a box with ice. To check the transfer Ponceau staining is performed on the nitrocellulose membrane and we checked the gel (12% SDS-PAGE) with Comassie Blue staining as well.
As you can see from the photos, there's a lot of protein that are still in the gel and it seems that we had an uneven transfer from the gel to the membrane (there are more HMW proteins than LMW proteins).
What could cause it? Maybe is it related with protein concentration? (there are 50 ug of proteins per lane). Is it possible to transfer all the proteins from the gel to the membrane?
The cell lysates were obtained with whole-cell method.