Science topic

Yeasts - Science topic

Yeasts are eukaryotic micro-organisms which belong to Ascomycota and the Basidiomycota phylogenic groups. 1,500 species have been currently described and are estimated to be only 1% of all fungal species. Most reproduce asexually by mitosis, and many do so by an asymmetric division process called budding.
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Hello all. I am repeatedly streaking one yeast strain (MATa MAL2-8C, SUC2) in 5-FOA plates aiming to disrupt the URA3 gene. After spreading thrice, the yeast is still able to grow in SD-URA plates. Am I doing something wrong? Do you guys have any tips?
It is my first time trying this widespread protocol to select URA3- cells. PS: I solubilized 5-FOA in DMSO, filtered and added to autoclaved medium (w/ YNB, SD w/aa, glucose 20%, agar 2%) to final concentration of 100mg/mL. Unfortunately I cannot try to disrupt URA3 by using CRISPR in the moment.
Thank you all.
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Do you add uracil to your plates containing 5-FOA (it doesn't seem so from your message)?
Also, are you sure your FOA plates are working? Do you have a positive control that you can add to the plates to test them?
I prepare my FOA plates by adding 0.1% 5-FOA in powder to hot agar (60-70 C) and stir with a stirrer bar then pour the plates (but I always check that the plates really work before using them!)
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Does yeast grow in MRS agar medium? If so, what might be the reason?
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@Drprabhurajeshwar C.
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Could anyone recommend a kit for total DNA (genomic and plasmid DNA) isolation from yeast cells? Thanks
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I do not recommend the kit. The zymolysis-based method is quite efficient with some modifications.
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I am using the PCR based methods to tag Saccharomyces Cerevisiae strains with antibody and fluorescent epitopes derived from plasmids. This method takes advantage of Yeast's homologous recombination system to integrate DNA fragments into the yeast genome.
Each plasmid region with the epitope also contains a selectable marker for transformant screenings (i.e. URA3/KANMX6). However, when amplifying tagging cassettes from these plasmids using primers with gene specific sequences homologous to the C-terminal regions of my desired genes (confirmed by gel electrophoresis), which then undergo DNA precipitation to be used as raw DNA for Lithium Acetate yeast transformations, only approximately 1/20 screened potential transformant colonies grown on selective media would have the desired edit in the gene I am working with after genomic DNA extraction, diagnostic PCR and agarose gel electrophoresis confirmation.
I have tried to extend the gene specific primer regions to theoretically improve homologous recombination efficiency for efficient transformations, yet my yield of success has been very low.
I am working with a mutant that has been shown to have Homologous recombination hindered to prefer Non Homologous End Joining. However, even in my wild-type strain, it is very difficult to have efficient and successful gene editing in my experience. Also, my negative controls (competent yeast cells lacking the tagging cassete with the selectable marker) also do not have any colony growth as expected so I know my transformations are at least visibly successful.
Has anyone who has used PCR mediated tagging methods found ways to improve transformation efficiency?
Here are the Papers I am using to aid in protocol:
Longtine, M. S., McKenzie, A., 3rd, Demarini, D. J., Shah, N. G., Wach, A., Brachat, A., Philippsen, P., & Pringle, J. R. (1998). Additional modules for versatile and economical PCR-based gene deletion and modification in Saccharomyces cerevisiae. Yeast (Chichester, England), 14(10), 953–961. https://doi.org/10.1002/(SICI)1097-0061(199807)14:10<953::AID-YEA293>3.0.CO;2-U
Lee, S., Lim, W. A., & Thorn, K. S. (2013). Improved blue, green, and red fluorescent protein tagging vectors for S. cerevisiae. PloS one, 8(7), e67902. https://doi.org/10.1371/journal.pone.0067902
Please let me know any tips for improvement!
Thank you!
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Hello,
TE buffer may work fine because commercial elution buffers contain Tris, but EDTA may cause issues during transformation. I am not completely sure about it but you can try water elution as a comparison to your method.
Colony PCR is truly inconsistent and sometimes specific transformation requires standardisation of the protocol. I use step-up PCR where I start the PCR with an annealing temperature 5C lower than the suggested annealing temperature, increase the annealing temperature 0.5C in each cycle for 20 cycles (so after 20 cycles, the annealing temperature = starting annealing temperature + 10 C, or, suggested annealing temperature + 5C), followed by running the PCR with the suggested annealing temperature for 10 cycle.
To prepare the template DNA for the colony PCR, I pick up a small amount of colony and resuspend in 5 micro litre of water (if dilution is required, then dilute 10 times), denature the colony at 95C for 10 min before adding the PCR mastermix.
Also it depends the type of transformation and the sequence variance at the C-terminal (3' of the gene). The gene deletion is tricky sometimes. For fluorescent protein labelling, I check the colony first under fluorescent microscope and use those shiny ones for colony verification.
Best wishes!
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How do you guys count your bacterial or yeast colonies? My fungus grows like yeast with distinct colony dots. I usually rely on the Fiji app(ImageJ) to manually tag and count them. What’s your go to method? 🧫 #colonycount #fungi
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Hi,
if I am interested in colony count, I just count them manually directly on the plate. But I do it only for "fun" after transformation of E. coli, so I don't need any picture or exact number in case I would miss something or so.
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I am working on yeast surface display library. After I work with yeasts for a while, I obviously see them at very bad conditions.
If I let the media stand for 1 min, I see cells aggregate very fast, form weird turbid media, and precipitate to the bottom super fast.
I assume it might be because the freezing with DMSO was harmful? But I honestly followed every step from a published protocol, so what could be the reason and what should I do?
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The DMSO should be fine, unless you are putting tons of it, but still it shouldn't cause aggregation. What do you have in your media? There are some chemicals known to trigger yeast flocculation, or maybe the genetic background of your strain or your expressed protein does it.
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We have been performing transformation associated recombination (TAR) in yeast. We believe that some of our yeast colonies (derived from a single cell) are maintaining two different TAR plasmids, one containing a large piece of DNA, around 35KB, the result of a successful recombination event into the plasmid. The other plasmid is empty, because it did not successfully receive a recombination event. It should be noted that the TAR plasmid contains the CEN/ARS origin of replication for maintenance in the yeast cell. The plasmid has a trp1 gene for selection on dropout medium.
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Hi!
Yes, Saccharomyces cerevisiae can definitely maintain two different plasmids in one cell. This is common in molecular biology experiments, where multiple plasmids with different markers (like TRP1) are used. As long as the plasmids have replication origins (like CEN/ARS), the yeast can replicate and pass them on during cell division.
In your case, it's possible for yeast to keep both a recombinant plasmid with a 35KB insert and an empty one, as long as the right selection pressure is applied.
I hope it helps.
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Hi all, I plant to creat a yeast mutant with two gene knockout. I want to use homologous recombination method. I found lots people will knock out one gene first, then continue with the second one. Could I knock out these two genes with kanMX and natMX at same time, and screen the positive colonies in media containg G418 and nourseothricin?Thanks.
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I agree with Michael J. Benedik, the sequential KOs will be far easier to obtain. Also, won't you want the singles as comparisons in your assays?
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I've tried DMSO before, but it doesn't work with the high concentrations of beta glucan
if you have any suggestions, I'll be happy to know it
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Thank you, sir, I'll try it
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I am conducting an experiment on comparative gene expression in yeasts using the real-time qPCR method. However, I am experiencing amplification in all my negative controls. I use MilliQ water, have changed my plate and adhesive setup, work with micropipettes dedicated exclusively to qPCR, wear gloves, and perform my work in a cabinet where I turn on UV light for a few minutes before starting.
Does anyone have any tips on where this contamination might be coming from or how to resolve it?
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Thank you for your effort and help. You gave me a lot of great tips that I will put into consideration and practice.
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Hello, colleagues.
I am currently working on the production and scaling of biosurfactant produced by Ustilago maydis. The methodology we currently use involves production in YEPS medium (1% yeast extract, 2% casein peptone, and 2% sucrose). We perform the extraction of MELs with a 1:1 ethyl acetate solution in the cell-free medium, and we recover the supernatant through evaporation. Additionally, we have searched the literature for other extraction and purification methods without much success.
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Is this toolkit only an extension for YTK (MoClo Yeast toolkit), does it assume that I have this kit and is based on YTK level 0 fragments?
Are there any regular YTK promoters or only the newly added inducible promoters?
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I think, newly added inducible promoter
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I've been working on fungal projects and recently, my petri dish has been constantly contaminated by 'this' thing. I'm not 100 percent sure if it is bacteria or yeast. Someone please tell me what that is and how to prevent it. Thank you so much!
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On the agar plates you have mites feeding on fungal spores and hyphae. The visible meandering patterns are the paths along which the mites move and spread the contamination.
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My cells can not survive with these bright spots (20x and 40x)!! I thought it was the yeast but now I don’t think so…. Please help me!
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The bright spots you’re seeing are likely contaminants, might be bacteria, fungal spores, or cellular debris.
In this case my suggestion is that:
1. do Gram staining or bacterial fluorescent stain to check if it’s bacterial or else?
2. Change to fresh sterile culture media to rule out contaminated supplies.
3. Assess cell viability just to ensure the issue isn’t due to the cell stress or death?
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This is a primary glial culture (taken from pups @PD2). Picture taken at 10DIV. Astrocyte layer is completely confluent. We are actually after the microglial layer underneath the astrocytes (Will use Saura et al. mild trypsinization procedure at DIV15). I'm thinking that the smaller cells might be yeast, except some of them have small projections. Thoughts? We are very strict about our cell culture protocols - all culture work is done in a BSC that is rigourously sanitized with 70%ETOH. Anything brought into the hood is sprayed/wiped with 70% ETOH, so not sure how yeast would get into this culture? The only thing I can think of is that during initial harvest, we don't have a downdraft table - it is done in the cell culture room on a counter that is wiped down with 70% ETOH and all instruments are soaked/dried prior to use. Could it be coming in via initial harvest?
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I think this is contamination of other cells because yeast is seen as different from this.
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why am i getting to see growth of yeast colonies after performing dilution spotting on the respective dropout media when only the empty prey and bait vectors were co-transformed? the yeast strain used was AH109 and the vectors used are PGADC1 and PGBDUC1
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I had done dilution series only starting from 10^0 to 10^-5. And the OD of the cells were 0.2
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Hey, i am working on oral probiotics, using MRS agar for lactic acid bacteria isolation, but now getting yeast contamination with bacterial colonies. how to prevent yeast growth on MRS agar without affecting bacteria?
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9/19/24
Dear Shubham,
The suggestion to use an anti-microbial is a good one. Yeast are eukaryotes. They can be inhibited by cycloheximide.
I hope this information helps you.
Bill Colonna Dept. Food Science & Human Nutrition, Iowa State University, Ames, Iowa, USA wcolonna@iastate.edu
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I have prepared a liquid broth containing Yeast (S. Cerevisiae) that i need to add from it to fermentation media of lignocellulosic hydrolysate.
I need to know based on what parameters do we add milliliters of yeast to fermentation broth?
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There is not a stright answer. When adding yeast to a mixture that's made from breaking down plant material, there are a few important things to think about. These include how much yeast you add, how much sugar is in the mixture, how acidic it is, how well the yeast turns sugar into alcohol, and how much stuff might be in there that could stop the yeast from working.If you get these things just right, the yeast will do a better job at making alcohol, which means you'll get more out of it in the end.
For most purposes, using a starting amount of yeast that's 5 to 10% of the total mixture is enough, especially if you're only interested in the final product and the stuff you're using to grow the yeast is the same as what you'll use to make the final product. Still it's important to consider the specific conditions of your fermentation. Factors like the type of yeast, what you're using to make the mixture, and how quickly you want the fermentation to happen can influence the best amount of yeast to use. Sometimes, you might need to adjust the percentage to get the best results.
However, if you want to study how the yeast grows and changes throughout the whole process, you should measure how much yeast you start with so you can keep track of how it grows.
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Good day to everyone,
I have little experience with culturing yeast, so I was wondering if someone could help with an issue I've encountered. I haven't seen many defined colonies from my plates of baker's yeast grown on agar, but I have been seeing these cloudy and spreading colonies of growth, and I am questioning if these were still yeast. Non of my control plates seem to have this issue, only certain treatement plates (the pink-tinted plate is also agar, just with some coloring).
Could this be contamination, perhaps from bacteria or some other fungi?
I appreciate any help, as I would like to avoid this in the future!
Best regards.
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This appears to be a matter of aseptic technique. Yes there a defined colonies - your inoculum is too heavy and you've either contaminated the plate or the culture is not pure.
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I need to perform yeast cell transformation after Golden Gate assembly in a 96 well plate. The cassette will be excised from the vector by RE digestion before transformation and usually, we clean the DNA by precipitation before transferring it into the cells. However, we are trying to avoid this "in between" step and I'd like to know more about alternative and faster purification/enzyme deactivation methods. Or else, what are the chance of success in case I jump straight to the transformation step?
Any suggestion, tips or comment are very welcome.
Thank you :)
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PCR amplification of the cassette is not recommended because the single units contain functional genes and are connected by small homologous regions. Any SNP introduced by the polymerase could be disastrous therefore, additional sequencing would be required to confirm the sequence, which will be both expensive and time-consuming.
Diluting the samples is definitely a good option, but excessive dilution will also reduce the transformation efficiency. I think I will try to compare different transformation playing with DNA amount, RE enzymes deactivation and dilution.
Thanks
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I am working on yeast transformation of a calcium channel. I performed twice transformation and no colony was observed after 3 days. The transformation protocol works well before. Is this gene toxic to yeast for the calcium channel activity? If so, How to achieve the transformation?
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Hello,
You did not mention what kind of transformation you performed. Do you want to delete an yeast gene or introduce a gene from different species?
If it is the case that you want to delete an yeast gene, you can check the null mutant viability and phenotype from the SGD database. If it is an essential gene, you can think about conditional knockout.
Which method do you use for transformation? LiAc-ssDNA-PEG method works very well for any yeast transformation.
P.S. - Yeast transformation has nothing to do with colonialism. :D
Best wishes.
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Hello Everyone,
I've been working with THP-1 cells and recently noticed some blackish clusters in the differentiated cells after 48 hours of differentiation (I've attached images of these differentiated THPs). To differentiate the cells, I use PMA at 100 ng/ml for 24 hours, followed by a 24-hour rest period in PMA-free media. Initially, I suspected a possible yeast infection, but the clusters didn't grow, and surprisingly, they even reduced after 72 hours post-differentiation. To be cautious, I discarded that batch of cells and started fresh.
However, in the new vial I revived from frozen stock, I observed something similar—reddish-black clusters(images of the undifferentiated THPs taken on the day of revival are also attached).
Has anyone else encountered something similar with their THP-1 cells? Any insights or advice would be greatly appreciated.
Thank you!
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That looks like clustered dead cells to me. Usually, it happens when you seeded too many cells into well plate. When some cells fail to attach, the dead/floating cells cluster together. It should not affect your results though.
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I want to make some yeast extractions with a protocol that uses Isopropanol and NaOH.
So in the end i will have an azeotrope of Isopropanol and Water that comes from the cell and NaOH.
Can you recommend a protocol to recover the isopropanol - so it can be reused in later extractions again?
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to recover and reuse isopropanol from a yeast extraction protocol using NaOH, the key steps would be:
1-Neutralize the NaOH if present in high concentration, such as by adding an acid like hydrochloric acid to form a salt that can be filtered out.
2- Break the azeotropic mixture of isopropanol and water, either by adding a salt or using a desiccant like molecular sieves to selectively absorb the water.
3- Distill off the purified isopropanol, potentially using fractional distillation to improve the separation, so it can be reused in future extractions. While complete recovery may not be possible, these techniques can help maximize the reuse of the isopropanol solvent.
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The plasmid that was transformed originally is correct (sequenced). I am expressing different gene targets from the yeast genome to optimize my protein. When I sequence the incorrect size, it matches one of my gene targets (also from the yeast genome) , not the one I transformed and it happened with many targets (they all match with one of the gene targets from the genome so this is not an operational mistake. My colony PCR Primers are on the promoter and the terminator and I check all the targets with the same primers. I got many right sizes too (targets from the yeast genome) in the same batch of transformation.
I have change primers and the Tms too, but the gel is either empty or the sequence matches with another target not the one I transformed.
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Aqsa Ahmed hello, i agree with Barbara Brezna as i also did not understand your question completely. Moreover, as much as i can understand, you are saying that when you checked your transformed vector via sequencing you get the correct target gene. However, in colony PCR the size is different but sequencing it revealed that it contains the sequence of your target gene. Now, there can be two possibilities
1. your plasmid contains the target gene but your insert (Target) is bigger than you are expecting.
2. Your colony PCR primers are amplifying a region which is bigger than your target gene.
It will be very helpful if you try and explain your question a bit more.
Thankyou
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hello everyone, do you have any advice on advice for Agar disk-diffusion yeast method?
like, medium agar to use and how many microlitres to use to wet the disk-diffusion?
thank you all for your help
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Agar diffusion/zone of inhibition will not be mkre in this case than qualitative. You can use any nonselective medium that supports growth.
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Image attached is at 20X. Cultured media for checking presence of bacteria and yeast and both tests were negative.
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looking like erythrocytes
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Hello, community,
Could you please clarify whether current legislation permits the reuse of algae biomass after it has been used to treat non-hazardous decontaminated laboratory organic waste?
Specifically, I want to understand any regulatory constraints or guidelines that might apply to this process, even if they are not directly concerned with using algae but other biological means (bacteria, yeast).
Additionally, are there particular conditions under which this reuse would be allowed or prohibited?
Cheers,
Gabriele
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It depends on the type of use because the treatments change the composite and the nature of the algae unless used as dead mass, which can only be used in commercial uses in the base of recycling.
It can never be used in research unless it is continuous research for previous research.
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I'm planning to perform BSA degradation using YCB and agar, but I need a protocol. Does anyone have one they'd be willing to share?
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Okay I will try, Thanks Phil
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Hi,
I've recently come across the colony of Geothrichum silvicola (based on sequencing result) isolated from soil with slimy morphology, unlike the colony morphology I've found on the journal below, which is moldy-like. Does anyone have experience with the difference morphology of yeast? and is there any chance the colony morphology of the yeast can change under certain circumstances?
I hope I have been able to convey what I want to ask.
Thank you for your answers.
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Bhavya G. thank you for the reference links!
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Hi all,
We performed Yeast-Cell-Glucose-Uptake assay to estimate the antidiabetic potential of the test compound.
The Metformin treated yeast cells, resulted in taking up more glucse than normal cells, however, the yeast cells treated with the test compound was observed to contain more glucose than normal yeast cells ?
Has anyone faced this similar problem ? Does it happen that the yeast cells will release glucose into the system than absorbing it.
the reaction was for 1hr in room temperature.
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Hello,
It's intriguing to hear that you've noticed similar observations in your experiments with yeast glucose uptake. Regarding the use of Metformin, it's curious that it doesn't enhance glucose absorption in yeast, as would typically be expected based on its effects in mammalian systems. It seems you've considered the possibility of glucose being synthesized by the yeast and subsequently released into the medium. Your approach to overcoming this by using higher concentrations of yeast in the experiment to avoid an increase in glucose levels in the medium sounds quite logical. This adjustment appears to provide a clearer understanding of the test compound's effects under controlled experimental conditions.
Could you share if you've found any practical solutions or additional modifications that effectively address these questions in your assays? I'm eager to learn from your experiences.
Thank you for sharing your insights, and I wish you continued success in your research.
Best regards.
Djihad CHENNA,
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I am already using BL21(DE3)pLysS cells but I am still getting leaky expression in pre-expression samples before IPTG is added. LB broth is tryptone, salt and yeast.
Thanks :)
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Add to 0.2% glucose.
Glucose has to be filtered sterilized not autoclaved.
If this does not work, you may want to reduced the plasmid copy number by introducing the pncB mutation (described by Jon Beckwith).
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?K
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French press, including Microfluidizer or any other mechanical high-pressure lyser, also glass beads (bead beating), manually for small volumes or with the help of a machine (BeadBeater). Some people use nitrogen grinding too, after freezing the cells in liquid nitrogen. Also, heating up the cells (like a colony PCR) may be enough to lyse them and get their DNA
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Is the instant dry yeast, or Brewer's yeast, Saccharomyces cerevisiae, had the ability to provide calcium and nitrogen minerals in addition to ethahnol ? Please support answers with published research.
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What are you suggesting is the source? Could you conduct chemical analysis?
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The yeast K. marxianus is well known for its ability to ferment de disacharide lactose. It is also able to ferment the disaccharide maltose or not? If you can share citations with respect to this issue I will be grateful.
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On the other hand, in this older work: Caballero, R., Olguín, P., Cruz-Guerrero, A., Gallardo, F., García-Garibay, M., & Gómez-Ruiz, L. (1995). Evaluation of Kluyveromyces marxianus as baker's yeast. Food Research International, 28(1), 37-41. it is stated "In spite of the fact that
K. marxianus is unable to ferment maltose (van der
Walt & Johannsen, 1984),...)
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Please ask if you need additional information.
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It would be useful to know what concentration of DAPI you used and fore how long in order to troubleshot.
Additionally, there are some very bright spots present, and I'm uncertain whether they are cells or debris. If you have exposure set on automatic this could be causing your microscope to adjust gain or exposure time to this very bright sports leaving your nuclei underexposed. Could also be that you have your contrast set in automatic in the software in which you are seeing/exporting the images and again this bright spots are interfering. If the second is true, then you would only need to adjust your contrast before exporting, overexposing those dots but correctly exposing your cells.
Hope this helps!
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I am new to Nanoformulations. I have done the synthesis of NLC for an antifungal compound using the protocol published in At the end of the process, I got a mixture of milky white solution with solid coagulants. Is this a correct form of how NLC looks? I have also doubts about %w/w calculations. Kindly help me with the calculation
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Dear Sathiyamoorthy,
According to my experience with NLCs, its final appearance depend on its composition, e.g. solid and liquid lipids, surfactants, etc.
My NLCs also get this "milky" appearance and I've noticed that It does not relate to particle's size and polydispersion index (PDI) by comparing to other NLCs with different composition. My colleagues recommend not to base it on the final appearance of the formulation unless it is something very unless it is something very out of the ordinary (such as phase separation).
I don't know about the solid coagulants though...Maybe the components are not sufficiently miscible?
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Just revived a sw480 cells from a 2019 batch.
Was normal on the first 2 days. After 5 day, the culture media became cloudy, and lot of suspicious white dots.
Is this yeast contamination, as I have never encountered this before.
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Yes, it is yeast contamination. At 100x magnification you will find yeast cells as separate round or ovoid particles or in chains of two to four or more particles and can sometimes be multi-branched. The appearance of chains is due to the most common form of replication called budding. You will find that yeast cells are larger than bacterial cells but smaller than the mammalian cells.
In early stages of contamination, you will not observe any change in growth media. But as the infection increases, the culture media will turn cloudy.
For more information on yeast contamination you may want to refer to the link below.
Best.
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I'm currently trying to capture a biosynthetic gene cluster using Transformation-Associated Recombination (TAR) in yeast. After identifying positive yeast clones, I extract, via an alkaline lysis method, the plasmid from yeast and electroporate it into E. coli DH10B.
However, I have not been able to find any positive hits upon cPCR on Eci clones despite testing about 100 colonies using the same diagnostic primers I used to identify the yeast positive clones. Then, on some of the E coli that I pick and miniprep, I consistently only see my original capture vector (with no gene cluster inserted).
The issue may lie in the yeast plasmid extraction, the transformation, or the plasmid isolation prep from E coli. I know that yeast plasmid extractions are hardly ever clean and tend to be "dirty," contaminated with yeast gDNA and other DNA it has inside due to the IPA precipitation required to perform the method. That tends to lead to poor transformation efficiency into E coli. But I feel like I'm stuck going in circles trying to bring the plasmid into E coli and isolating it. if anyone has ever worked with TAR and has experienced any troubleshooting at this stage of the process, I would appreciate insight. Or any advice on things I can try to better improve yeast plasmid extraction or transformation/isolation in E coli. Many thanks!
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Transforming and isolating a large plasmid in E. coli can sometimes be challenging, but there are several troubleshooting steps you can take to optimize your protocol:
  1. Quality of Plasmid DNA: Ensure that the plasmid DNA you are using for transformation is of high quality and free from contaminants. Purify the plasmid using a reliable method such as commercial kits or CsCl gradient centrifugation.
  2. Transformation Efficiency: Optimize the transformation efficiency by using competent E. coli cells that are prepared fresh and are highly competent. You can prepare your own competent cells or purchase commercially available ones.
  3. Transformation Protocol Optimization: Review your transformation protocol and ensure that it includes proper heat shock conditions, incubation times, and recovery steps. Optimizing these parameters can significantly improve transformation efficiency.
  4. Plasmid Copy Number: Large plasmids may have lower copy numbers compared to smaller ones, making them harder to isolate. Consider using bacterial strains that are optimized for maintaining large plasmids, such as certain E. coli strains like DH10B or DH5α.
  5. Selection Markers: Ensure that your plasmid contains appropriate antibiotic resistance genes for selection. Use antibiotics at the correct concentrations to select for transformed colonies while suppressing growth of non-transformed cells.
  6. Screening for Transformants: After transformation, streak the transformed cells on selective agar plates containing the appropriate antibiotic. Incubate the plates overnight at the optimal temperature for E. coli growth.
  7. Isolation of Large Plasmids: Large plasmids may require special isolation techniques to prevent DNA shearing during purification. Consider using methods such as alkaline lysis followed by cesium chloride (CsCl) gradient centrifugation or commercial kits designed for isolating large plasmids.
  8. Verification: Once you have isolated the plasmid DNA, verify its size and integrity by running it on an agarose gel or using other analytical techniques such as restriction digestion or sequencing.
By carefully optimizing each step of the transformation and isolation process and troubleshooting any issues that arise, you can increase your chances of successfully transforming and isolating large plasmids in E. coli.
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I tried glycine buffer but the yeast cells acidify the buffer so the pH goes down to 7-6 overnight. I need something that will stay around 9 and that isn't toxic to the cells.
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To maintain a high pH (pH 9) for yeast cell suspensions, you can use a buffer that is effective in this pH range. Tris buffer is commonly used for maintaining alkaline pH and may be suitable for your needs. Here's how you can prepare Tris buffer at pH 9:
  1. Tris Base (Tris(hydroxymethyl)aminomethane): Dissolve Tris base in distilled water to make a 1 M stock solution. The molecular weight of Tris base is 121.14 g/mol, so to make a 1 M solution, dissolve 121.14 g of Tris base in 1 liter of water.
  2. Adjust pH: Adjust the pH of the Tris solution to 9 using concentrated hydrochloric acid (HCl) or concentrated sodium hydroxide (NaOH). Use a pH meter or pH strips to monitor and adjust the pH as necessary.
  3. Final Dilution: Once the desired pH is achieved, dilute the Tris solution to the desired concentration for your experiment. Common working concentrations for Tris buffer range from 10 mM to 100 mM.
  4. Sterilization: Filter-sterilize the Tris buffer using a 0.22 μm membrane filter to remove any particulate matter and microorganisms.
  5. Storage: Store the Tris buffer at room temperature (if using within a few weeks) or at 4°C for longer-term storage. Avoid repeated freeze-thaw cycles.
Tris buffer is widely used in biological research and is generally compatible with yeast cell suspensions.
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Hello!
When constructing the plasmid, I used the same promoter. However, the direction of each expression cassette promoter in the obtained plasmid is the same, and the plasmid with the opposite direction of promoter cannot be obtained. In this case, I am worried that homologous recombination within the plasmid may occur after integration into yeast, resulting in the loss of some expression cassettes. How should I do toget the plasmid with the opposite promoter direction?
Thanks in advance!
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To reverse the direction of two identical promoters in the construction of a plasmid, you will need to manipulate the DNA sequence of the plasmid. Here's a general guide on how you might accomplish this:
1. Identify the promoter sequences.
2. Design PCR primers to amplify the region containing the promoters in the reverse orientation.
3. Perform PCR to amplify the desired region.
4. Purify the PCR product.
5. Clone the purified PCR product into a suitable vector.
6. Verify the orientation of the promoters in the newly constructed plasmid.
7. Transform the plasmid into a suitable host organism.
8. Conduct functional assays to confirm the activity of the reversed promoters.
These steps involve manipulating the DNA sequence through PCR, cloning, and transformation techniques to achieve the desired orientation of the promoters in the plasmid.
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I would like to make solid plates without the metals iron, copper or zinc for growth studies in yeast. I'd like to test the effects of the absence of each of these metals individually while leaving the other two constant. Does anyone have a method for doing this? Normal YPD contains trace amounts of all these metals. I'd like to make something similar to what is found here, but in solid media.
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Investigate potential of agarose or silica gel matrices. You'll also need to control water used in prepration and of inoculum.
To what level of detection will you consider "absence"?
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Good morning,
I used TCA/aceton protocol to precipitate proteins. I incubated vesicles from yeast with TCA. After centrifugation 14000xg, 10min at 4°C I saw black pellets in each samples... What could happen? Is there any option to purification such protein pellets?
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Thank you all for your answers!
I will try to wash again my pellets.
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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
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You may also try 3000-4000 rpm for 5 minutes. This keeps the cells alive and it works for our experiments.
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Most of the GFP-ATG8 studies use yeast-derived plasmids rather than integrating into the genome. For the ones that integrate GFP-ATG8 into the genome, they are using a commercially available URA3 marker for the URA3 locus, which is not available in BY4741. Is there any other way than making a new plasmid? Thank you!
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Hello Songlin Wu
GFP tagged Atg8 is stable when it is tagged at the N-terminal. You can prepare a PCR cassette with Atg8 N- and C-terminal homology following GFP sequence in frame. For the selection of the right transformation, you need a selection marker which can be either an auxotrophic marker (for yeast strain) or antibiotic marker (Kanamycin/Nourseothricin). BY4741 strain has uracil auxotrophy (ura3Δ 0). This URA3 gene will be integrated first on the 5'-UTR of the gene, but then you have to eliminate it using Cre-recombinase.
Best wishes.
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I'm running RNA extracted from Saccharomyces cerevisiae in a Tapestation 4200. The RINe value is excellent, but the sizes of the 18S/28S bands are lower than expected: ~1000 and ~1800 instead of ~2000 and ~4000, respectively. The internal control (lower marker, 25nt) in each sample ran as expected. Yeast don't seem to have a hidden break in the rRNA.
Has anyone experienced a similar problem?
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Hi Shon, you're right. The 28/18 ratio is great, but the positions seem to be off. Are you sure the lather you used was not for DNA by any chance?
Plus, may I ask what was your purification method? the product is really clean.
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Hello everyone. I am working on antimicrobial properties of yeast isolated from kombucha. So I chose 2 (out of 4) yeast strains with the highest antimicrobial properties, ran a PCR test and then send my samples in order to be sequenced. The BLAST results showed me Candida parapsilosis. I wonder, is this a relevant finding? I searched and read a few articles about this yeast strain's role in fermented foods, it's potential as a probiotic (somehow?) and it's technological properties in fermentation, especially in fermenting glucose and even findings of Candida spp. in kombucha. I am currently writing my thesis, yet as a Master's student, I am somehow worried and not sure to even report it. I am quite sure of my PCR test, as I saw sharp bonds on my electrophoresis gel and used proper primers (ITS and NL-4). Is this data considered relevant or am I being overly-nervous?
Thanks a lot in advance!
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yes bit score
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Iam trying to optimize the Comet assay protocol for Yeast cells in my lab but iam unable to lyse the yeast cells with the regular protocol and hence those cells are seen intact even after the treatment. I looked upon some protocols which suggested the use of Zymolyase/ Lyticase for the Yeast cell wall lysis, but my lab doesnt have either of these enzymes and hence can someone suggest any other enzymes apart frombthe above mentioned that I can use for the same? Also can I use Lysozyme for digesting the Yeast cell wall?
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Thank you so much W.J. Colonna and Malcolm Nobre for your responses.
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Yeast from a small scale fermentation was heat inactivated and stored in a refrigerator after centrifugation - about 60-70% water content.
The Yeast was planned for lipid/sterol extraction. Sadly i got sick for about a month before i could do the extraction.
Now the refrigerated sample is moldy.
Do you know what influence mold can have on such a sample - if it may be still usable after removing the mold, what the mold may have used/altered as food source to grow?
As to get to this point was quite time consuming I would prefer not to need to repeat the experiment to this point - and maybe find a way to save the sample.
I found a paper investigating the Effect of bug damage and mold contamination on fatty acids and sterols of hazelnut oil (DOI: 10.1007/s00217-016-2778-x)- where the lipids/sterol decreased. Not sure if this is similar in a liquid solution with yeast settled at the bottom.
Do you have any experience and/or advice for such a situation?
Any tip and help is much appreciated
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Well, you immediately have a problem with reproducibility if you use the samples, so it depends a lot on what you need the lipid for and what types of conclusions you would be making. If you are going to make claims about the fatty acid profile of the yeast, well the month long storage itself may have changed that in addition to whatever changes the fungus has caused. Other factors include which fungus (or fungi) are contaminating the sample, what residual growth substances were in the liquid between the yeast cells (e.g. were the cells washed before storing or are there still medium components present). How much lysis of the yeast occurred during the storage time? If all you need is some microbial lipid and you don't care what sort of profile it has and are not going to make conclusions about the productivity of the yeast it could be okay to extract your cells (might as well just leave the fungus with the yeast and extract everything). Separating the fungus from the yeast would be very challenging. I would expect the total lipid content to have gone down. Otherwise, unfortunately, you need to start over and hopefully utilising what you have learned in the first round, things go more efficiently this time.
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What solvent should I use for the yeast cells? Precautions to take when preparing the solution (pH...)? concentration range for non-toxic efficacy?
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I prepare in deionizated water at the concentration of 600mM, because the solubility is near it (I think it is 660mM) and then I filter the solution to sterilize, because I think it degrades in autoclave. I use the final concentration of 20mM and it is not toxic and improves the growth in oxidative stress conditions, but other people already saw this effect at less concentrations, like 10mM.
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Hi everyone!
I would like to start researching the response of human mast cells to yeast. Which commercial human mast cell line do you think is the best? I have read that LAD-2 is difficult to passaging and ect. What do you think about HMC1.2 and LUVA?
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The two known human mast cell lines, HMC-1 and LAD2, are derived from patients with mast cell leukemias. The HMC-1 cell line represents very immature transformed cells and they contain no chymase, very little tryptase, and do not express functional IgE receptors.
LAD2 cells are functional mast cells but have a widely variable doubling time, an unstable phenotype in culture, and can be difficult to freeze and recover.
On the other hand, LUVA cells respond to IgE receptor cross-linkage and resemble mature human mast cells functionally and morphologically. They do not exhibit autophosphorylation of the c-kit receptor and respond to recombinant human c-kit ligand, the stem cell factor, with accelerated proliferation. However, neither the stem cell factor nor any other exogenous growth factor is necessary to maintain the survival or proliferation of LUVA cells. LUVA cells are the first mast cell line derived from a patient with no clinical mast cell disorder and no mutation in the c-kit receptor. They should be valuable for studies of human mast cell function.
I recommend the use of LUVA cell line for your study.
You may want to refer to the articles attached below for more information.
Best.
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Hello.
I have a question about these two behaviors, the Warburg and Crabtree effects. I am a master's student and my teacher said to me that Warburg occurs in cancer cells and Crabtree in yeast, basically that the Warburg effect on cancer is analogous to the Crabtree effect on yeast. However, I see some scientific articles that say that cancer cells have both. In yeast, I understand that high concentrations of glucose inhibit oxidative respiration. I am confused. Can someone explain this to me? Cancer cells only exhibit the Warburg effect or also exhibit the crabtree effect?
Thanks
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The “Warburg effect” is defined as an increase in the rate of glucose uptake and preferential production of lactate, even in the presence of oxygen. The normal differentiated cells will rely primarily on mitochondrial oxidative phosphorylation to generate the energy needed for cellular processes. But cancer cells instead will rely on aerobic glycolysis, a phenomenon which is termed as “Warburg effect”.
Aerobic glycolysis is an inefficient way to generate ATP, however, the advantage it confers on cancer cells has been unclear. But there is a thought stating that cancer cells acquire and metabolize nutrients in a manner conducive to proliferation rather than efficient ATP production, and the associated mutations acquired by cancer cells enable them to function in this manner.
In some cancer cells, sometimes, depending on the absence or the presence of glucose and the environmental conditions, the cancer cells can reversibly switch between fermentation and oxidative metabolism. This short-term and reversible event is referred to as the “Crabtree effect”. This reversible shift might be an advantage to cancer cells, as it would allow them to adapt their metabolism to the rather heterogeneous microenvironments in malignant solid overgrowths.
So, cancer cells show both the effects, depending on the environmental conditions, they can switch reversibly.
Your teacher is right when he/she said that the "Crabtree" occurs in yeast. Some yeast species such as S. cerevisiae use fermentation even in the presence of oxygen when glucose concentrations are sufficiently high. The use of fermentation in the presence of oxygen and at high glucose concentrations is referred to as the “Crabtree effect”. Yeasts that display a “Crabtree effect” are Crabtree-positive while yeasts that do not display a “Crabtree effect” are Crabtree-negative.
You may want to refer to the article attached below for more information on “Crabtree effect” in yeast.
So, when high concentrations of glucose or fructose are added to the culture medium, respiration is frequently inhibited. This phenomenon of “Crabtree effect” is observed in numerous cell types, particularly you will find this effect in proliferating cells, which would include not only tumor cells but also yeast.
Hope this clears your doubt!
Best.
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I am relatively new to conducting Western blots. I employed the alkaline lysis method to extract whole proteins from yeast. Subsequently, I introduced the Opti Protein Marker (ABM, CAT log NO: G252) and completed the gel electrophoresis. The proteins were then transferred to a nitrocellulose membrane using the wet transfer method. Following the transfer, I performed a Ponceau staining, during which the protein markers were clearly visible.
Moving forward, I proceeded to block the membrane for 1 hour in 5% BSA in TBST (Initially, I attempted blocking with 5% non-fat skimmed milk, but encountered high background signals). Subsequent to blocking, I washed the membrane with TBST (3 x 7 mins) and TBS (1 x 5 min). Following this, I incubated the membrane overnight at 4 degrees Celsius with a primary antibody (Beta-tubulin, Rabbit IgG Polyclonal), 1:1000 dilution in 5% BSA in TBST). Afterward, I repeated the washing steps with TBST (3 x 7 mins) and TBS (1 x 5 min).
For the next step, I incubated the membrane with a secondary antibody (Rabbit anti-Goat IgG (H+L), HRP, Polyclonal, 1:10,000 diluted with 5% non-fat skimmed milk in TBST) for 3 hours. Subsequently, I performed additional washes with TBST (3 x 7 mins) and TBS (1 x 5 min). Finally, I carried out chemiluminescence detection with an exposure time of 30 seconds.
However, my results were unsatisfactory as I observed multiple nonspecific bands, and the protein marker disappeared. I seek assistance from experienced researchers, especially those familiar with Western blotting of yeast proteins. I would appreciate any insights or suggestions to identify and resolve the issues.
Additionally, please refrain from suggesting the use of monoclonal antibodies, as it is currently beyond my budget constraints.
Thank you in advance for your help.
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If you or others have not tested this antibody before then it's possible this is just a "bad" antibody that binds to multiple proteins. It's very common for antibodies to show different results than what is advertised by the company selling them.
I can't find much information about your protein marker, but you should make sure this marker is compatible with chemiluminescence. Ponceau staining will stain all proteins, so that's why you could see the marker protein there. But if the protein marker is not modified or your secondary antibody does not bind to the marker proteins, then you will get no chemiluminescence signal from the marker protein. You can read more about this at the bottom of this page: https://www.thermofisher.com/us/en/home/life-science/protein-biology/protein-biology-learning-center/protein-biology-resource-library/pierce-protein-methods/chemiluminescent-western-blotting.html
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LAPT broth: 1.5% Bacto Peptone, 1% Bacto Tryptone, 1% yeast extract, 1% glucose and 0.1% Tween 80
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Thanks Robert, but the LAPT media I am using doesn't contain lactose.
LAPT broth: 1.5% Bacto Peptone, 1% Bacto Tryptone, 1% yeast extract, 1% glucose and 0.1% Tween 80
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I have a query about carabcillin antibiotics is good option to put it in ypd agar for growing yeast or is it possible to grow them without antibiotic?
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If you've a pure culture, there's no need for antibiotics. For isolation of a yeast like fungus from mixed culture, carabcillin (assume carbenicillin) is ok but reportedly not so great vs. Gram + bacteria.
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HELP!
Our lab has recently discovered these dancing blobs in our TC cultures - across cell lines, primary cells and organoids. So far they are not behaving like any bacteria/fungi/yeast we have ever come across (not responding to antibiotics/antifungals and no turbid media). They seem to be amorphous and both extra/intracellular..
Someone has suggested they may be protozoa? If anyone has seen something similar or is an expert microbiologist please help us identify them!!
(Picture included for attention but really need to watch the videos to distinguish from cells/debri)
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Looks like amorphous debris.
Have you cultured for microorganisms?
btw - Kingdom Fungus "yeasts"
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I've been trying to isolate a low copy (CEN/ARS) plasmid from yeast that has selective markers for both yeast and bacteria (Trp and Amp). However, when I try to recover the plasmid I can't get any positive bacterial clones when transforming the DNA. Running the DNA on a gel it looks like I mostly isolate genomic DNA.
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I've had success with the Zymo prep protocol. You'll need to purchase the zymolase enzyme, as it is needed to lyse the cell walls. If your plasmid is on the large size, try warming your elution buffer prior to use and incubate it for 1 minute prior to elution.
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Does anyone have experience with or can provide references to studies that have employed the RNA Plant Micro Kit for RNA extraction from yeast cells?
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Yes You can. Even the kit says the same can be applied for fungal extraction
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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!
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I haven't done it, but I found this publication by J. P. Aris where they describe a method to isolate yeast nuclei:
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I was just thinking could it be possible just because 2- 5 hr storage of my yeast cells before soptting could switch to anaerobic or post diauxic shift
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Sorry but your question is a bit confusing. The title of your question is asking about plates, but the text of your question seems to imply storing the cultures before plating on plates.
But there is adequate aeration on agar plate surfaces, at least until the colonies get large (there is oxygen deficient in the center of large colonies). If you are storing a culture in a tube prior to spotting then there is of course no aeration. Whether or not they switch to anaerobic growth depends on what they are stored in. If in neutral buffer then not much changes. If in medium then they might switch metabolic states. However this is not really of concern because you are then plating them on on plates in aerobic conditions so by a few generations they will have converted to aerobic metabolism.
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I am studying the phagocytic activity of frog blood cells. After incubating frog whole blood with yeast cells, I had this microscopic image; Could be possible for frog erythrocytes to engulf yeast cell?
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I have disrupted yeast gene in genome now I want to verify it without delaying it for overnight culture and then genome isolation so can I directly check it (gene disruption )from transformed yeast colony plate by colony pcr?
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You can do a quick check by colony PCR. If you want to publish your findings you'll need to do a more thorough test (sequencing).
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Kind and warm greetings to all
I'm working on Antifreeze proteins(AFP) that is expressed by Yeast So which method \ assay is more reliable and effective to detect AFP by western blot or ELISA with references ? and why? Kindly if anyone working now or previously on Antifreeze protein can you guide me.
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ELISA (Enzyme-Linked Immunosorbent Assay):
  • Advantages: ELISA is highly sensitive and can provide quantitative data about the concentration of the anti-freeze proteins. It is a widely used method for quantitative protein analysis.
  • Limitations: ELISA requires specific antibodies and may not be able to differentiate between different isoforms or modifications of the anti-freeze proteins.
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The media that was used YCB?
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Thank you Dominique for your reply.
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Protocol used for transformation (TAKARRA)
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I am not able to see any colonies during single drop out experiment and I can't figure out what went wrong that I'm not getting any colonies?
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Upon linearization of plasmid DNA for electroporation mediated transformation in yeast, is it necessary to precipitate out the DNA from the restriction reaction via alcohol (With 75% ethanol). Isn't it possible to just deactivate the restriction enzyme via heat treatment and proceed towards transformation using the same restriction mixture?
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In addition to what Robert Adolf Brinzer writes, adding some carrier such as tRNA helps to precipitate your DNA if it is at low levels.
Regarding using ethanol, it works but you need a final concentration of around 75%, in other words adding 3 volumes of 100% ethanol to your solution.
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I did yeast two hybrid transformation a few days ago and no yeast colonies grow in the DDO+ 225 AbA plates after 3-5 days. OD600, carrier DNA, competent cell, as well as the amount of AD and BD plasmid were correct according to TAKARA manual. Can anyone know the reason why no colonies could grow in the plate?
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Try giving it a few more days. Sometimes you'll need 7-10 to see some growth. Also, try making single plasmid selection plates to check that both your bait and prey plasmids are giving good transformation and growth on their own.
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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
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Robert Adolf Brinzer Dear Robert I can use this NEB DNase for my purpose.
Could you please take a look at this product and let me know your idea? Already bought one DNase but was not good enough to do this experiment. Thanks in advance
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Hello,
I'm working on an optimization experiment related to yeast growth and the best conditions for growth. Optical density is a measure of yeast growth, and from the literature, this yeast strain grows best at 30 degrees Celcius. Suprisingly the results have shown that optical density is higher at a lower temperature, around 20 degrees. What statistical test should I use if the null hypothesis is that the highest optical density is achieved at a temperature of 30 degrees Celcius?
Please be kind. I have minimal background in stats; I'm just a lab tech. Thank you
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If you want to check if difference in optical density is statistically significant, you can use simple unpaired t Student test. Your null hypothesis is OD at 20 degrees Celsius == OD at30 degrees Celsius. The alternative hypothesis is OD at 20 degrees Celsius != OD at 30 degrees Celsius. You prepare samples at 20 and 30 degrees, measure OD thrice, calculate means and standard deviations, and you perform t Student test.
If you want to check how OD changes in function of temperature and express it in equation, you can prepare samples at 20, 25 and 30 degrees Celsius, measure OD of each sample thrice, calculate means and standard deviations and finally try to fit the function, perhaps linear or quadratic.
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So far I have seen that organic acids are used to kill off spent yeast from the brewing industry before being fed to animals such as pigs. The amount of acid needed to achieve a satisfactory kill off seems to be quite cost prohibitive so I was wondering why I have never seen formaldehyde containing products being used for the control of yeast in animal feed in regions where formaldehyde is allowed? Thanks
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Typical commodity pig feed in general does not need to be sterilized unless there are certain animal products in it. Acidifying the brewers yeast halts its growth which can be problematic during processing and storage, especially if it is mixed with fermentable sugars such as might be found in various grains. Also, some brewers feel their strains are highly proprietary and would want to prevent them from being easily cultured.
Formaldehyde might not be an allowable feed additive. As Martin noted, it would reduce its nutritional value, likely by forming undesirable adducts with the proteins.
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We are working with Pichia pastori and must take our yeast to another country. What are the considerations for the transportation? I mean, besides the air company considerations.
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Hello, I send yeast every week abroad. Are they alive or dried? You will need some Polystyrene Thermocol Box, maybe some Icepack, and always check with customs! Invoice on the outside of the package (in a transparent sleeve) with the customs declaration, etc. Most countries have postal agreements with each other, but some have their own rules when it comes to yeast. If they are alive, you need to ship in less than 24 hours. Hope that can help.
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Dear ResearchGate Community
Which type of yeast does exist in this field?
Is there still a possibility of yeast contamination despite the continuous use of 70% ethanol and Bleach 10 % ?
What can be done if such contamination is observed?
Thank you in advance for your time and consideration.
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Fungal spores are not inactivated by 70% ethanol. You need to use bleach or other strong disinfectant.
Could be a Candida species of yeast, although the photo is rather poor.
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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?
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Disaggregating yeast chains can be challenging, especially when working with certain species like Saccharomyces cerevisiae that tend to form large cell aggregates. Here are some techniques you can try to improve yeast cell separation:
  1. Enzymatic treatment: Enzymes like zymolyase, lyticase, or mutanolysin can break down the cell walls and release individual cells from the aggregate. You can add these enzymes directly to the cell suspension and incubate for 15-30 minutes before washing and resuspending the cells. Be careful not to overdo the enzymatic treatment, as excessive exposure can damage or kill the cells.
  2. Sonication: This method uses sound waves to disrupt cell clusters. Briefly expose the cell suspension to ultrasonic energy using a probe sonicator or bath sonicator. This will break apart the cell aggregates into smaller clumps. Immediately after sonication, transfer the cells to a fresh tube and proceed with your desired analysis. Take note that sonication may also cause cell damage, so use caution and monitor cell viability.
  3. Mechanical disruption: Applying physical stress through vortexing, pipetting, or passage through narrow tubes can help break up cell clusters. Gently vortex the cell suspension for short periods, then immediately stop and check for cell separation under a microscope. Repeat this step several times until the desired level of disaggregation is achieved. Alternatively, pass the cell suspension through a narrow gauge needle or capillary tube to physically disrupt the cell aggregates.
  4. Chemical treatment: Some chemical agents like sodium dodecyl sulfate (SDS), Triton X-100, or saponin can solubilize lipids and break down cell membranes, leading to the release of individual cells. However, these agents can be harsh and require careful optimization to avoid cell death or altered cell properties. Start with low concentrations (e.g., 0.01-0.1%) and gradually increase them while monitoring cell viability and separation efficiency.
  5. Hypotonic shock: Cells exposed to hypotonic solutions (low osmolarity) experience water uptake, which can swell and eventually burst cell membranes, releasing individual cells. Prepare a solution with distilled water containing 10-20 mM glucose, 10-20 mM HEPES (pH 7.4), and 0.1-0.5 mM EDTA. Add this solution to the cell suspension and incubate for 10-15 minutes at room temperature. Then, rinse the cells with PBS and analyze them promptly.
  6. Flow cytometry pretreatment: If you plan to analyze the cells using flow cytometry, consider performing the above steps just before running the samples through the flow cytometer. This ensures minimal loss of cells due to sedimentation or aggregation during the analysis.
  7. Combinatorial approaches: Try combining multiple methods mentioned above for improved disaggregation efficiency. For example, first treat the cells with enzymes, followed by brief sonication, and finally, passage through a narrow tube or addition of a hypotonic solution.
  8. Yeast genetic manipulation: Consider engineering the yeast strain itself to express surface proteins that facilitate cell separation. One approach could be to introduce genes encoding for adhesion molecules with defined epitopes, allowing you to target and separate individual cells using specific antibodies.
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