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Bacteriophage - Science topic

Research and developments in the field of bacteriophage related to its classification, Molecular biology, Bioinformatics, etc. and their biochemical and genetic interactions with bacterial communities and co-evolution.Virusoids, viriods, prions, bacteriophages and other virus like particles all have a very magnificent way of living their parasitic intracellular life and they do have a significant effect on the evolution of smallest and the largest living organism on earth.
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Hello,
My colleague carried out an experiment in which she grew epithelial cells in a well plate followed by infection with Pseudomonas and treatment with phages. She wanted to determine the effect of the phages on cfu over 2, 4 and 24hrs. At each time point she removed the supernatant from a well and diluted this ten fold from 10-1 to 10-8. These dilutions were plated onto a plain Columbia agar plate and incubated.
The results in my opinion are opposite to what would be expected e.g. the most bacteria appears in the most diluted sample and no bacteria is seen in the 10-1 dilution. My colleague thinks that it's possible that there were many active phages in the first dilution that continued to kill the bacteria and that in the lowest dilution the number of active phages is lower so the bacteria grew the most.
I cannot understand how the bacteria wouldn't be diluted out as well as the phage and wondered whether it's more likely to be a error in dilution or plating.
Thank you for your help!
Rosemary
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Hi Andrew,
Thank you for your reply to my question. I just have another couple of questions:
1. I didn't mention in my original message that a MOI of 1 was used for infection/treatment as our other experiments had shown the phage to perform best at this ratio. Does this make any difference or would the results still be reliant on the respective growth rates of the bacteria/phage and relative proportion of uninfected bacteria?
2. As you said in the first couple of dilutions there would still be phage capable of infecting growing colonies. However my understanding is that phages need a top agar medium or bacterial lawn plate upon which to survive. I know the bottom agar provides nutrients but how would the phage be able to continue to kill the bacteria once absorbed into the plate. Is it just that they are in relatively close contact in each drop of solution on the plate?
I hope these questions make sense. Thanks for your help!
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I have the phage sample titer and the bacteria colony forming units (CFU) - but how can I calculate how much of each I want to plate knowing I need to plate 1000x more phage than bacteria cells?
Ex: Bacteria host: 2.8E8 cfu/mL
Bacteriophage titer: 1.8E5 pfu/mL
What volume of host and phage do I plate knowing I need to plate 1000x more phage than bacteria cells?
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Are you really sure you need 1000X more phage than bacteria? That would be quite unusual for any experiment that I can think of. Also a titer of 1.8E5 pfu/ml is really low.
But given that you know the CFU or PFU per ml for both, it is just a simple math problem to do your calculations. WIth the numbers cited above, adding 2.8 ul of cells and 1.8ul of phage would give you 1/1000 phage per cell.
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I will be grateful if someone help me with the anti-tuberculosis assay particularly Luciferease Reporter Phage assay for my work.
I am available to discuss further.
Thank you in advance.
#tuberculosis research
#anti-tuberculosis
#anti-microbial screening
#TB
#TB research
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The National Institute for Research in Tuberculosis- Indian Council of Medical Research (ICMR), is an internationally recognized institution for Tuberculosis (TB) research, India. It is a Reference Laboratory and a WHO Collaborating Centre for TB doing anti-tuberculosis assay particularly Luciferase Reporter Phage assay
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Hello,
I have one dilemma how to avoid phage contamination of BSC cabinet via bacteriophages and father cross-contamination of all samples and lysate ?
as filters are suitable for bacteria ? not for phages !
Or it is better to work old style on bench with fire ?
Thanks
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I think that most air filters (such as HEPA filters) will not completely remove viruses and bacteriophages. The filter pore size is effective for removing microbes but not viruses (although it will reduce it to some degree). However if your BSC has UV lamps for sterilization, those are effective for killing any contaminating phages in your cabinet.
However I worked on phages just on the bench top for many years without any problem. So if you have good sterile technique then it should be fine either way.
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Many phage purification methods use ultracentrifugation and We do not have ultracentrifugation facility in our Institution. Therefore a versatile methodology with minimal loss of phage particles with high viability would be more useful for us. Kindly suggest me some procedures
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I see. I am afraid I don't know. But sometimes even simpler protocols work. If youre phage lysate are concentrated enough, maybe it works? If it is not too much work, then give it a try (and post the answer here).
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When I examined one-step growth curve experiments for bacteriophages, I mainly saw two experimental setups; one is based on bringing bacteria and phage together and precipitating after incubation and resuspending the pellet while the other is based on diluting after incubation and re-incubating that sample and continuing. After incubation of bacteria with phage, after centrifugation of the mixture, what is the purpose of resuspending the pellet again, forming a new mixture and taking a sample for DLA? I can't understand why the two protocol types have different paths.
Another thing is that I already bring bacteria and phage together and incubate in these experiments but I wasn't sure if I had to put bacteria back into it while making DLA the sample I took to create this one step curve.
Article example for two one step-growth curve protocols;
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Dear Michael J. Benedik , thank you for your valuable response and advice.
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Hi we performed a dilution series 1:10 of different amplifications of the same phage library in Xl-blue bacteria, and we see a strange pattern. In the first dilutions of the reamplified library there should be a matt of bacteria but we se few single colonies and then the spot gets progressivly thicker to then beahave like a normal titration. i include 2 pictures one if a library that behaves normally i.e a solid spot of bacteria that graduually gets to songle colonies the furtger down the dilution series and then one that starts off single colonies and increases to a solid spot and then go down.
Has anyone seen this pattern before? has any one any idea of what this can be caused of?
lytic phage infection should not look like this. that would produce lytic placs and just not very few colonies.
does it have anything to do with the helper phage? should very high concentration of phage/helperphage or something else inhibit bacterial growth?
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First of all I find this experimental protocol very confusing, I am not sure what you hope to learn from it. But the short answer is that if you have more phage than cells, you will have very reduced numbers (or none) colonies growing in the spot. But once the cells start to outnumber the phage, then you will get spots that look reasonably identical. You are not likely to ever see plaques doing it this way.
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Hi i am doing phage display and in the amplification stepr growing bacteria dubbelinfected with M13 mibrary and VCSM13 helper phage in 200 ml SB media (10g mops 20g yeast extract 30g tryptone / liter) is behaving strange. it has a orange/ pink color (see picture )and the pellet does not stick well to the wall of the centrifuge bottle (see picture) has anyone experienced this before? what is the reason and is it still fine?
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While I have no experience specifically culturing phage-containing bacteria, the pellet not sticking to the centrifuge tube often happens when the pellet itself becomes really slimy due to cell lysis, or more rarely, biofilm formation. If uninfected bacteria grown under the same conditions are doing fine, then it's likely not biofilm formation and it's your phage reproducing and causing cell lysis or extreme morphological changes.
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If crRNA sequences are ~20 nucleotides, the genome of each virus offers a multitude of potential crRNA sequences, yet natural CRISPR systems work remarkably well.
How do bacteria know which 17-23 nucleotide sequence to extract as crRNA?
Other questions: are crRNA sequences often off-target (i.e., incorrectly target non-viruses) or are they usually precise and specific to a given virus? Do bacteria extract multiple nucleotide sequences for the same viruses? Is there a pattern to how bacteria choose crRNA sequences?
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Hanna Alalam not related to this question, but you may find this recent study on phage defense interesting: https://twitter.com/Nitzan_T/status/1546588441319800833?s=20&t=s1E_IlkZEuzBmS3AcewPIQ
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During phage purification by plaque assay. At 3rd step of plaque (small plaque) purification my plates are showing smaller size as well as bigger size plaques. Can a single phage form different size plaques?
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Michael J. Benedik and Pierre Béguin thank you for your answers. I will proceed with my experiments keeping the given suggestions in mind.
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10-fold dilutions of phage (initial concentration 3E9 PFU/ml) + bacteria (1E5 CFU/ml) were run in triplicate on microplate. Plate was incubated overnite at 37oC, with OD600 measured every 30 min. Growth curves were plotted to determine active concentration of phage (see attached file). Looking for an explanation for the dilutions showing delayed growth only in 1 or 2 wells (dilutions -2 to -9, but especially in -2 to -7). Additionally, it is confusing that the pattern does not track with sample dilution (e.g., -2, -3, -4 dilutions). Bacterial resistance to phage is well established, but if that were the case wouldn’t it be expected that the replicates be consistent for a given dilution?
Any help is appreciated.
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If you are trying to enumerate phages then I think the traditional overlay method for titers or or spot titers is going to be much more repeatable and meaningful. If you do a full dilution series by spot titers you can get an actual count. I think monitoring growth of the host will prove to be very problematic.
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Hi, I'm working on a project where we're using S. aureus and corresponding bacteriophages. I did a spot plate and found out that my phage stock is roughly 2.0x10^7 PFU/ml. I want it to be at least 10^10 PFU/ml. My method for growing the bacteriophages has been to make agar plates and then use top agar that's been mixed with the bacteria and the phage dilution. How will I go about making more stock solution that is the appropriate concentration? Do I just have to make it and then test it?
Thank you.
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I agree with Dr. Michael J.Benedik answer
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I am studying about Staphylococcal phage and I obtained lytic bacteriophage. Now, I want to precipitate Bacteriophage using pEG6000 with 2.5 M NaCl. How can we preparation PEG-6000 20% + NaCl 2.5 M for bacteriophage precipitation? and how can we adjust their volume (?uL of peg6000/? uL of NaCl), Could you help me, please?
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how can I create drought conditions by using PEG 6000 for drought-resistant rhizobacteria in the laboratory to check their drought-resistant attributes
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I am trying to isolate DNA from bacteriophages specific to Klebsiella pneumoniae using the standard phenol-chloroform methodology. But the DNA yield/concentration is very low. Is there some alternative methodology specific to Klebsiella pneumoniae phages? Thanks in advance.
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Malavika B H Wow, yes 1013 is a very high titre. That's not the problem then. Not sure what else to suggest.
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Hi, researchers
I am looking for growing the dual-species biofilm, including proteus mirabilis and other UTIs causing pathogens and I will treat them with the proteus phages. Does anyone know any protocol or method to grow this dual-species biofilm where I can characterize this dual-species biofilm and monitor the proteus phage treatment to this dual-species biofilm? if anyone can help me it will be much appriciated.
thanks
Akash
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Darío Lago Espartero Thank you so much for the help but proteus is a very terrible pathogen, it behaves differently and it overtakes the other species
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What is the time required for bacillus culture to reach log phage?
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You may also try reducing the percentage of top agar
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Hi, Iinserted gene of Reverse transcriptase into pPLc245 plazmid. If I'm right, this plazmid doesn't code cI857 represor. For expression od reverse transcriptase from this plasmid I used E. coli DH10B. RT was expressed after increase of cultivation temperature to 37 °C. Before this thermo induction I cultivated E. coli DH10B with pPLc245 at 28 °C. But at this temperature RT was not expressed. Why is this possible? Is E. coli DH10B coding cI represor or pPLc245 could contain gene for this represor?
Thank you for all responses.
Bohuš
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If your plasmid really is pPLc245 (and not some derivative of it) then it should not carry the lambda repressor gene and you should get constitutive expression at any temperature. So I can't really explain your result unless RT does not fold properly to have activity at RT or 28.
I'm actually a bit surprised this worked at all for you though, generally plasmids with superstrong promoters like lambda pL are not very stable under conditions where there is no repressor.
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I am trying to construct a phage library via Kunkel mutagenesis. The template ss-DNA was prepared from phage following Sidhu's protocol. One primer containing the degenerate codon was used to perform Kunkel mutagenesis with 3:1 molar ratio added. However, after an overnight room temperature incubation, I didn't see the complete conversion of ss-DNA to ccc-DNA as described in the published protocol. I am wondering if anyone has ever seen this before? Any suggestions to optimize the reaction condition? Tricks?
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Thanks, Michael J. Benedik , I noticed that some people actually cut out the correct ccc-DNA from gel... I can certainly do that! And I was also trying to do a slow ramping method to anneal the long primer onto the template.
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Recently, I'm working on a bacteria isolated from soil. It's hard to transfom. I have used chemical transformation and electrotransformation methods. They all failed. Then I found phage transduction. But I found little articles which used phage as a tool to transfer plasmid. I want to know why? If I want to use phage as a tool to transfer plasmid, is it feasible? I think maybe the reason is it's hard to isolate a lysogenic phage.
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It is possible to use some lysogenic phages to transduce plasmids but the real answer is very complicated. First it will depend upon the phage, some phages such as lambda will only package their own DNA (or DNA with the cos site), so you can package a cosmid for example but not a typical plasmid. Also many (most?) phage package specific amounts of DNA that fit the phage capsid. For example P1 phage packages around 90kb and is not going to effectively carry a small plasmid, or one much larger than that.
So there may be packing restrictions based on site and there may be size restrictions. Having said that, there do exist phages that are more flexible. But you would need to know the underlying biology of the phage to know. Or if you have good selections you can try and see if it works or not.
Regarding conjugation, again your plasmid will have to have the appropriate transfer signals for the conjugative apparatus to recognize and transfer the DNA. If you are working with large natural plasmids they might carry their own system, but if these are laboratory engineered plasmids then you would need to look whether it has the right sequences or not.
Also, do you know for certain that the plasmid you are attempting to introduce into the new strain is compatible and will replicate in that strain?
Having said all that, it might be worthwhile to work at optimizing your electroporation method, it might be just some minor technical problems.
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I have sequenced my phage genome from Qiagen. I'm using Galaxy CPT for genome assembly and annotation. My Question is that How can I confirm that I have assembled the contigs of my phage correctly?
Given that i have selected contig having the greater length and a coverage of 1000. Then I did blast on NCBI and e value of first selected sequences 35 is 0.0
Please help me out.
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Hello Saleha,
There are different alternatives to evaluate the assembly using bioinformatics tools in general.
1- QUAST:
You can get a basic metrics of the assembly using this tool. In the results, you can see the how many contigs, N50 value, which is a quality metric in terms of contigs length in your assembly, the longest contig length and so on. However, N50 is like mean or median value so I would say N50 would not give you a powerful idea of the assembly.
2- BUSCOs (Benchmarking Universal Single-Copy Orthologs):
You can analyze your assembly in evolutionary core gene sets in different lineages. Basically, every genome must have some orthologous genes. You can see how many orthologous genes and how many of them are duplicated and fragmented in BUSCOs results. As you go down taxonomically, the number of genes in gene sets will of course increase.
3- REAPR (without reference):
It evaluates your genome assembly in terms of bases. You can see mis-assemblies, errors, error-free bases.
4- Reference genomes:
If you have somehow references or closely related genomes, you can mapped your genome against reference or closely related genome and have an idea how it looks.
As you can see, there is no only one way to assess your assembly if it is correct or not. More, those alternatives cannot say if your assembly is 100% correct or not. So, you can focus primarily on the compactness, in other words, less fragmented (you can use QUAST, and BUSCOs) then you can run REAPR.
Hope that helps. You can read following articles for more information.
Best,
References:
1- Gurevich A, Saveliev V, Vyahhi N, Tesler G (2013) QUAST: Quality assessment tool for genome assemblies.
2- Liao X, Li M, Zou Y, et al (2019) Current challenges and solutions of de novo assembly
3- Simão FA, Waterhouse RM, Ioannidis P, et al (2015) BUSCO: Assessing genome assembly and annotation completeness with single-copy orthologs
4- Hunt, M. et al. (2013) ‘REAPR: A universal tool for genome assembly evaluation’, Genome Biology.
5- Earl, D. et al. (2011) ‘Assemblathon 1: A competitive assessment of de novo short read assembly methods’
6- Bradnam, K. R. et al. (2013) ‘Assemblathon 2: Evaluating de novo methods of genome assembly in three vertebrate species’, GigaScience.
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I spiked some clam tissue with stock phage and recovered 2.6E6 cp/uL using digital PCR.
here are all my #s:
digital PCR - 2.6E6 cp/uL
amount used in dPCR reaction- 5uL
Amount eluted for extraction- 50uL
Amount used for extraction- 400uL (phage stock)
is this correct?
2.6E6 cp/uL x 50uL = 1.3E8 cp / 400uL = 3.25E5 cp/uL
thanks!
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Can you clarify if the Digital PCR value of 2.6E6 cp/µL is the concentration readout from the dPCR instrument? Because I'm pretty sure that is higher than the detectable range for dPCR.
The standard way to get to concentration of phage in the stock before extraction would be as follows:
  • Concentration readout from dPCR (cp/µL) * volume of dPCR (µL) = amount of template (cp)
  • Amount of template (cp) / volume of extract added to dPCR (µL) = concentration of template in extract (cp/µL)
  • Concentration of template in extract (cp/µL) * volume of eluent (µL) = amount of template extracted (cp)
  • Amount of template extracted (cp) * recovery (%) = amount of template spiked (cp)
  • Amount of template spiked (cp) / volume of spike (µL) = concentration of spike (cp/µL)
You don't mention knowing the recovery, so assuming that it is 100% (spoilers: it won't be) and your dPCR reaction volume is 20 µL (a guess assuming you are using a Bio-Rad QX200) equations look like this:
  • 2.6E6 cp/µL * 20 µL = 5.2E7 cp total template in PCR
  • 5.2E7 / 5 µL = 1.0E7 cp/µL in the extract
  • 1.0E7 * 5.2E8 cp extracted
  • 5.2E8 * 1.0 = 5.2E8 cp spiked
  • 5.2E8 cp / 400 µL = 1.3E6 cp/µL in the original spiked sample
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Recently, phages are used against the MDR pathogen in humans to combat the diseases. Wouldn’t our immune system try to eliminate those bacteriophages? How these techniques are successful?
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Phages have been used in human therapy for many many years, so this is not a new approach. As to how they escape our immune system, first it will depend upon the mode of delivery, for example if administered orally it won't really interact in any significant way with our immune system, so that should be fine. It might be more of a problem were the phages to be delivered by injection into the circulatory system. But more relevant is that the phages are likely to have acted very rapidly (hours) whereas an immune response will take days.
It may well be true that for some bacteriophages they would only be effective one time if the body did mount an immune response, but again this would primarily only be relevant for phage in the circulatory system.
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  • I prepared grids by adding 10 μL bacteriophages (109 pfu/ml) and allowed them to dry on air. I used 1 ml 2% uranyl acetate solution and put grids in it. Then, waited for 1 hour but I did not get any image for my phages.. Could anyone help me?
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Negative staining is a very old (about 70 years) but still useful method for routine virus visualization. It is very well described in many textbooks, papers, etc. Personally, I prefer this way:
1. Put a droplet (10ul) of viral suspension on parafilm;
2. Put a grid (ultrathin formvar, stabilized with about 1nm of carbon) on the droplet (face side down) and incubate for about 30sec;
3. Pick the grid up with a sharp tweezer, then gently dry it up with filter paper from the edge, and then immediately put the grid (face side down) to a droplet of 1% UA (water solution). Incubate from 20 to 60 sec (depending on the virus), then pick the grid with a sharp tweezer and gently dry it up with filter paper from the edge.
It's necessary to remember that the suspension should be as pure as possible and the film on the grid should be very thin.
Good luck!
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Other than uranyl acetate, are there any negative stains that can be used for phage electron microscopy?
Is the visualization better if uranyl acetate is used?
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PTA has been used, as Indranil has suggested, but so has uranyl acetate. You will need to look up a good method so you have a good working method.
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How to measure the density of bacteriophages from a soil suspension like we measure the density of bacterial cells? I would like to know the phage density in the soil suspension to decide the inoculate density. Thanks in advance.
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The problem with trying to determine phage numbers in a sample is that it will be entirely dependent on the host you use. So you can enumerate E. coli phages on E. coli, or Pseudomonas phages on Pseudomonas etc. But really the only thing that is going to work is to titer by doing serial dilutions and plaque counts on the host that you are interested in using.
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Hi guys,
Im looking to understand the mechanism of which resistance could develop against phage. Research has shown that phages can elicit the production of antibodies. And I was thinking that following secondary treatment these levels may be produced at a higher level.
How exactly would this occur, is it due to memory cells b or t and how would this lead to an increase in antibody production
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Some bacteriophages of Xylella  fastidiosa, Liberibacter spp., Spiroplasma spp. could offer protection against the plant disease. Although several published experiments show some effects in reducing symptoms development, the tested control measures are not able to completely eliminate the bacteria from diseased plants. What is the future of phage-based control of tree diseases caused by these bacteria?
EFSA Panel on Plant Health (EFSA PLH Panel), Bragard C, Dehnen‐Schmutz K, Di Serio F, Gonthier P, Jacques MA, Jaques Miret JA, Justesen AF, MacLeod A, Magnusson CS, Milonas P. Effectiveness of in planta control measures for Xylella fastidiosa. Efsa Journal. 2019 May;17(5):e05666.
De Leon, Victoria S. Investigation of'Candidatus Liberibacter Asiaticus' Prophages in Texas and Florida. Diss. Texas A&M University-Kingsville, 2020.
Chipman PR, Agbandje-McKenna M, Renaudin J, Baker TS, McKenna R. Structural analysis of the Spiroplasma virus, SpV4: implications for evolutionary variation to obtain host diversity among the Microviridae. Structure. 1998 Feb 15;6(2):135-45.
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Characterization of Novel Virulent Broad-Host-Range Phages of Xylella fastidiosa and Xanthomonas
Authors: Stephen J. Ahern, Mayukh Das, Tushar Suvra Bhowmick, Ry Young, Carlos F. GonzalezAUTHORS INFO & AFFILIATIONS
Best Regard.
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If I understand the Lambda genome correctly it seems that some of the sequence of genes isn't the most logical. When the phage enters the lytic cycle its late rightward promoter first transcribes the genes that actually achieve host lysis including the lysin, spanin, and holins (genes R, S, RZ) and only then continue through to the structural and DNA packaging genes (genes nu1 to J). Wouldn't it seem more logical for the phage particles to begin being produced before cell lysis is initiated allowing a bit of a head start to release the most phage as possible? Is it that the difference in expression time between the gene products is really inconsequential, or does the actual lysis require a buildup of the lysins, etc that allows sufficient time for phage production? What would be the effect of engineering the phage to reverse the order of the genes?
Also regarding the capsid proteins, it is known that the stabilizing decoration protein gpD only attaches to the capsid after it is formed. Yet it would appear from the sequence of the genes that gpD is actually expressed before the major capsid protein gpE. Again is the order of gene expression just not as significant as I would have thought or is there a reason for the way the genome is arranged?
Thanks in advance for helping with what might be a silly question.
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One assumption you are making that is not correct is that cell lysis does not actually begin upon the expression of the lysis proteins. The actual lysis event is carefully timed and occurs much after their expression is initiated (about 40 minutes for lambda) and this timing is inherent to the lysis proteins themselves. So there is not actually an issue here.
It is hard to say whether the gene order is significant or not in an evolutionary context. There is clearly a constraint on genes for related functions being generally grouped together, this is probably so they are inherited as a unit upon recombination events, but beyond that I don't know.
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Hi guys,
I am currently writing my dissertation on the phagemid based delivery of CRISPR.
I'm currently looking at the limitations, specifically now looking at the immune response against phage. However, I'm struggling to see how phage is actually administered and where the immune response would occur.
Is the main route of transmission through the bloodstream, where antibodies and macrophages act on the phage and form a response or does this not occur?
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Hello Ben,
Intravenous administration of phage therapy started in the 1940s.
Kindly read article titled " Safety and efficacy of phage therapy via the intravenous route" DOI: 10.1093/femsle/fnv242.
Furthermore, topical application also demonstrated no side effects, see the attached article
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I am currently working on a spot assay for a bacteriophage that infects Ralstonia solanacearum. There are 3 types of stock bacteriophages that I use:
1. Original stock isolated from 2016, I think without the addition of glycerol
2. Stock of bacteriophage - ralstonia solanacearum which has been filtered using a syringe filter 0.2 and stored without glycerol (2021)
3. Stock of bacteriophage from plaque stored in a mixture of ralstonia solanacearum : glycerol 40% (1 : 1). This plaque is from stock restorations 1 and 2 (also 2021)
all three stocks were stored at -80. The results of the spot test, stocks with glycerol (number 3) showed a clear zone/plaque, other stocks did not. I doubt whether the clear zone that appears is plaque or because the growth of bacteria is inhibited by glycerol. Please help with the information if anyone has experience with this. Thanks.
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Glycerol can cause osmotic stress on the bacteria leading to clearings on your bacterial lawn if too concentrated.
You can try to produce stocks with varying concentrations of glycerol (as long as phages remain stable) and dilute with SM buffer to reduce glycerol concentration before use. Phage titre will also be reduced.
Note that phage tolerance to freeze varies and you may need to try a different approach.
I'll leave this article for other options:
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I do not know here is phage or not. I incubate over night at 30oC. Vibrio parahaemolyticus causing AHPND and spotting phage on agar.
Please give me advices
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I see two issues on your plate. First of all you do not have a lawn on the plate so you won't see plaques without a lawn.
Secondly your "phage stocks" are not sterile and still have a lot of cells in them.
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I have to perform therapeutic experiment of phages in chicken model. I am confused about dose calculation. could not find any specific formula to calculate it.
Is there any specific formula to calculate phage dose related to animal body weight or any other?
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Also a lot will depend on phage characteristics. Phages with high burst rate, good killing properties, short latent period and less BIMs will work even in low concentrations and low dosage. Phage therapy is "state of art".
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Hello everybody,
I wanted to isolate phages against Micrococcus luteus from human nose swabs. I tested nose and mouth swabs in a spot assay on M. luteus for phages and the results were clear spots for both. The swabs were diluted in sterile saline only.
Then I cultivated 10ml M. luteus in TSB media to OD600 of 0.5 and added 100µl filtered swab material to the culture and added 100µl of saline to another culture as negative control.
I incubated both at 37°C shaking. The next day the M luteus with swab sample was clear and the negative was opaque, at OD600 of 2.0.
For me, this is a sign of lysed bacteria probably due to phages lysing them.
So, I took 1 ml of the whole culture (not centrifuged) and filtered it through 0.2µm and added it to a new culture, incubated overnight shaken at 37°C. I also tested the filtrate in a new spot test.
The spot test as well as the culture did not show any signs of lysis or spots.
So, I think there might be something other lysing agents in human samples, like lysozyme or something. Or the phages are inactivated fast or go temperent... Does anyone have an idea, what this result could mean?
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If the filtrate from the "lysed" culture is not causing lysis in the next round, then most likely you do not have a phage. Even if it were a temperate phage that lysogenized, you would still have phage present. But if it were a temperate phage, then the lysogenized cells would grow and the swab culture would be dense.
You might just do a simple dilution of the swab solution and see if you don't get lysis with a 10 or 100-fold dilution then you may just be diluting out some chemical or proteinaceous agent. You could test to see if the "lysis" effect goes away with heating or proteinase treatment to distinguish between the two.
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Hi Phage fam!
I am currently doing some research on phage DNA density regarding a particular phage we are studying; I am trying to calculate the DNA density as bp/nm^3, and i have the capsid volume and the originally deposited DNA sequence of our phage, but it's a head-full packaging phage and I wasn't sure if a "complete" genome included the coding sequence of one genomic concatamer, or if it included redundant DNA as well? I considered trying to figure it out from the sequence but it's not exactly my area of expertise, so if anyone knows i'd be very grateful!
P.S. If anyone knows if %redundant DNA can be assessed from sequence, i'd be very curious to hear about that too :)
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Normally DNA sequencing is occurring on a population of molecules and not just a single molecule. Therefore the "average" of all the sequences should give you a single genome sequence without redundancies. However, as the others have pointed out, this is something you can quickly analyze from the sequence itself (and it might be annotated).
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Does anyone know a good way of predicting E. coli K-antigen identity from NGS data? It seems like there are lots of options for O and H antigens (e.g. the excellent ECTyper) but none for the K antigen. I'm hoping to type some strains to better understand the host ranges of a phage collection. Any help gratefully received!
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If you have assembled genomes, this should be easily done. The most simple way I could think of right now is similarity based analysis, for example, BLAST.
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I have assessed the effect of trypsin and pepsin on phages. Thus, I want to compare the normal concentration range with the concentration where phages remain viable.
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I am so grateful for the data. it has been of great use
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I am a third-course microbiology student and I am researching relationships between cyanobacteria and bacteriophages that infect them. For my project, I need to do an alignment of host genomes, and all the programs and websites that I used say that my file is too big. 180-120 MB is there any program, that could do an alignment because neither MEGA or UGENE could do it( was waiting >24 hours and no results). I don't have any knowledge in programming and I have windows operating system. So maybe someone has a solution? Thanks in advance.
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Why do you want to align multiple genomes? What is your aim?
Ask this question to your course leader and understand the concepts of your the analysis you want to perform. Those tool you mention and the tool recommended in above post are not meant to align genomes.
Useless processing without proper understanding of aim would not lead you anywhere.
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The lab I work in is looking to isolate phages from water sources, but we have had no luck in the past. What is the best protocol for isolating phages from water samples? We work primarily with Mycobacterium hosts.
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Filter the water using filter paper to separate coarse materials. Then, consider doing serial dilutions to isolate bacteria on LB agar plate and purify the bacteria to obtain pure colonies.
To isolate phages, try spot test method. Mix bacteria culture together with the water sample and LB, centrifuge and filter using 0.22 μm. Next, do the standard plating by transfer 100 μL of the filtrate and 100 μL of the bacterial culture onto 1/2 LB agar plate, pour 3 mL of Top agar on top of it (you can adjust the quantity where applicable) and incubate overnight. Observe the clear zones of phages on the next day. All the best!
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I am isolating a bacteriophage specific for Staphylococcus aureus bacteria from stool samples. I used overnight bacterial culture and NA medium. I pre-incubated for 1 hour for double agar. Why are my zones irregular as in the picture?
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Your top agar is not well dissolved.
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I'm having trouble getting clear images of my bacteriophage under TEM. The process I've followed thus far is described below. Any help would be appreciated.
I've isolated bacteriophage active against Staphylococcus aureus bacteria from wastewater.
I grew these to high titer by infecting liquid cultures (S. aureus grown to OD600=0.1) with a low dose of bacteriophage (Multiplicity of infection = 0.1 to 0.05) and incubated until lysis was observed ~3-5 hours later.
These lysates were spun down at 5000g for 5 minutes and filtered through 0.22µm filters.
Chloroform was added (0.1 volumes / 1ml in 10ml), mixed, and left to incubate at room temp for 15-20 minutes with inversions every so often.
These were then centrifuged at 3220g for 10 minutes and care was taken to remove the lysate without touching the cell debris and chloroform layers.
We then used ultrafiltration columns (Cytiva: Vivaspoin 20, 100,000 MWCO) to resuspend the phages (trapped on the column membrane) in pure SM-buffer.
30µl of high titer phage was then fixed using paraformaldehyde and sent for TEM.
TEM images are stained with Uranyl acetate.
Please advise.
Thanks, Josh.
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I agree with Michael J. Benedik . Just a thought: it looks like the darkened area is in the region of the highest phage aggregation. Maybe it is the reason for higher charging? Please note, that the resolution in that region is also affected - drastically reduced.
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I did phage isolation specific for Campylobacter jejuni/coli. I use sample that derived from chicken's feces which suspended in SM buffer and store at 4 C for 24 hours, After that filtrated the sample by 0.2 uM pore size filter. 10 ul of filtered sample was dropped on bacterial lawn media (0.4% w/v agarose). The plates was further incubated at 37 C for 48 hours.
Finally I got result as shown in figure.
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It looks to me that you have phage all over the lawn (all the little plaques everywhere) suggesting that your host strain or some thing you used to make the top agar layer had phage contamination. Then the large spots also look like they could be phage as well, although it could also be anything that is inhibitory to bacterial growth.
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I'm isolating bacteriophages from the faeces sample, but I can't see clear zones. I observe bacterial growth within the phage zones on the double agar plate that I left overnight.
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Bacterial growth occur on phage plate due to strain specifity and O.D culture.
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Hello, I work with recombinant proteins (endolysins) derived from lytic bacteriophages. This week I am doing some tests with my recombinant protein and I noticed that it has lost its lytic activity. My transformed cells (BL21(DE3)) are stored at -60°C in glycerol. Do you know if this temperature can be harmful to the expression of my proteins?
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Loss of expression and activity of your protein is possible only when one is growing an undesired clone (either a contaminant or gene-deficient clone). I completely agree with Sir Pierre Béguin. Since endolysins are proteins that are toxic to bacterial cells and you are trying to express them in an E. coli host that shows leaky expression. It, therefore, becomes important to regulate this leaky expression by changing your E. coli host. Good luck!
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Dear all, I want to introduce random mutation to double-stranded DNA phages using the Cobalt60 gamma source. I searched for a suitable dose rate that is high enough to mutate the phages but not too high to damage the phage particle integrity. I found one paper (Bertram 1992) that used 60 Gy/min. Does anyone have any suggestions or documents that I can refer to? Thank you very much.
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That makes sense. Thank you very much.
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Since temperate phages would not produce bacteria lysis, thus it can't be used in phage typing. So how to determine the hosts for temperate phages?
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Pierre Béguin is correct that temperate phages still produce plaques. Therefore determining the host for a temperate phage is no different than determining a host for a lytic phage, does it form plaques (or lyse) the host.
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This is a fundamental question about basic phage library construction. I know that people can construct library by either phage vector (eg. M13KE) or phagemid+helperphage. Based on my understanding, phage vector has everything we need to assemble a phage and we insert our library pieces into the vector; xhagemid is a plasmid only express pIII protein with library pieces; and helperphage contains everything on wild type phage. So why we cannot use a blank M13KE vector as a helperphage? and if we only need a pIII protein source, can M13KE with library pieces insertion serve the same function as phagemid and produce monovalent library by adding additional helper phage?
thank you
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Thank you!
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Dear Community,
I would like to know in phage display library construction, after transforming host cells (e.g. E. Coli SS320) with phagemid and helper phage (M13K07) and overnight culturing to collect the phage particles in medium, can we save the host cells by frozen for re-culturing in future ? The reason is because the construction of antibody library is tedious and expensive, it would be great that can be able to re-culture the host cells and re-collect the phage particles for future selection. Thanks.
hon
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The bias issues comes from the fact that some phages grow faster or more robustly than others due to the insert.
I don't have any data that would make me discern whether keeping infected cells frozen or reinfecting with phage would be better. Maybe someone else on here can answer that.
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The titer of Phage stock 2.2* 108 was added in MOI 5. After 5 minutes, the remaining phage count was 3.7* 106 . What is the degree of phage absorption?
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hi, Hope this will help you.
Isolation and characteristics of new phage JK004 and application to control Cronobacter sakazakii on material surfaces and powdered infant formula - ScienceDirect
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Other than sequencing to check for presence of integrases, how do we experimentally determine if a phage is lytic or lysogenic?
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First, most lysogenic phages will form turbid plaques (and not clear ones).
Secondly you can isolate the lysogens from the center of those turbid plaques and show; 1) they are immune to the same phage so that the phage will not form plaques on the strain; and 2) it will release phage again (usually). An overnight culture from such a presumed lysogen can be centrifuged to remove the cells and then you will find phage in the supernatant that you can titer.
There are always some exceptions to these but in general these are the simplest way to experimentally determine.
If you want a molecular approach you could develop PCR primers to the phage and grow presumptive lysogen and show they carry the phage. But to do this properly you need to restreak a few times and to make sure you don't have any contaminating phage along for the ride that would also give you PCR product.
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Can I check by Western Blot with the protein sample of the bacteria that was infected by the phage at different times? Does Western Blot Sensitive enough to check one of the phage proteins expressed in the host bacteria cell?
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iTRAQ-based Proteomics Analysis may help you at initial level.
Thankyou
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I try to isolate human phages against Staphylococcus capitis and Micrococcus luteus, both found on human skin. First, I take sterilized and moistened swabs (with 0,9% NaCl) and rub it on a defined surface area of 16cm^2 on the forearm, face and scalp. Then, I transfer the swabs to 5 ml of phage buffer, shake it vigorously and filter it through a 0.2µm membrane.
To test if there are phages I do the spot assay and dispense 3 drops of 5µl of the samples on a S capitis lawn.
I already tested bacteriophage spot assays of e coli and it worked well.
I think the phages of s capitis and m luteus on the human skin may be lysogenic. Can someone reccomend how to induce the lytic cycle or has someone already experience in isolating human skin phages?
Thank you!!
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Hope this paper will help you for isolation. Isolation and characterization of bacteriophages from the human skin microbiome that infect Staphylococcus epidermidis.
Thankyou
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i would like to know, how viral DNA's are unique when compared to that of the bacterial DNA. Why doesn't the CRISPR-Cas system activate for a bacterial DNA when it gets transferred to the bacteria through conjugation or transformation?
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I would like to know the detailed protocol, especially about Cas9-gRNA preparation.
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I have a question, I try to isolate phages from environmental samples, and during this process, I face with a clear glass tubes after overnight incubation (host+phage), but I spot nothing on the agar plate after titration process (bilayer agar method). after several days of amplification, I keep seeing the same results; clear glass tubes, no plaque on the agar. Do you have any idea what is happening?
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Michael J. Benedik unfortunately no picture with me now, but I will post it definitly in my upcoming manipulations. thanks for your recommendation.
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I am looking for a company that sequence phages. I need DNA extraction also. Any suggestion is welcome.
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Macrogene
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One of the tests performed to check the polyspecificity of an antibody selected in vivo or in vitro is to test it in ELISA against BVP extract. My idea is to start already during the selection campaign to add the BVP extract as a blocking reagent for my phage libraries.
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I usually use biotinylated-antigens in solution and magnetic beads for the actual selection, so I was wondering if I can add the BVP directly in the phage solution, but it is not too much work to preclear the phage before passing them through an ELISA plate well or an immunotube coated with BVP
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Hey, I have a small problem in the observation of virus plaque using LB broth, the problem is that the Top agar looks distributed around the plate without a confirmation which gives a bad result for plaque. Can anyone help, please? #top_agar #phage.
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It depends on your choice of host strain. You cannot be so sure that you have phage infection cum lysis if the phage is present and it is host specific or if the phage is absent. While you did not explain the method you used, if your agar was not given enough time to cool before pouring, it will prevent infectivity and plague formation.
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Hi,
I’ve just started to work with phage display. My goal is to construct a short peptide-displaying phage library. For this purpose I would like to use the phagemid which is composed of M13 phage DNA and use MCS to insert the degenerated oligonucleotides into the pVIII gene. Which phagemid vector and which E. coli strain will be the best choice in this case? Thank you in advance for your help.
Kate
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You might wish to consider purchasing a peptide display library that is already built, it is likely to save you a LOT of time. I know that NEB sells them and I'm sure other vendors do as well.
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I am using imageJ to quantify fluorescent bacteria and phages. Ihave tried and deciding to use either OTSU or adaptive tresholding. Is there a good system to determine what is the best strategy? Also once, the settings for adaptive tresholding have been set, should they vary between different slides or should they remain constant? What would be the best strategy to go about accurate quantification
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Thank you will do!!!
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there are numerous drugs for newly diagnosed, treatments are needed to enable repair and remyelination.
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thanks wolfgang, i've been considering probiotic strains previously
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What is the remarkable aspect of using phage promoters?
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I think the identification of regulatory elements, primarily promoters, which are specific DNA regions responsible for transcription initiation. Muskan Gupta
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Can you stain and image (via TEM) bacteriophage directly from lysate? (10^9 PFU/ml suspended in SM-buffer)
Or do phages need to be purified via other means prior to imaging?
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Joshua Iszatt,
Data below could be helpful:
How do you check methylation status?
Currently, there are three primary methods to identify and quantify DNA methylation. These are: sodium bisulfite conversion and sequencing, differential enzymatic cleavage of DNA, and affinity capture of methylated DNA (1). Restriction enzyme based differential cleavage of methylated DNA is locus-specific.
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i am trying to figure out how to propagate bacteriophage- phi 6.
i try different media (NBY, 2XYT, LB) and different tempearture (22-28oC)to propagate as mention in different article.
i am worried about the bacteriophage-phi 6 stability as well may be its not stable.
anyone having a experience to propagate RNA bacteriophages that might be helpful in my research.
need your suggestion thanks in advance
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Thanks. let me know if you learn anything new on phi6 propagation, Im still struggling with it.
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I am looking for some method for phage inactivation with out damage of bacteria, for sterility test of phage preparation by USP 71.
Thanks in advance
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I think I better understand. You have your phage cocktail and wish to inactivate it really as a negative control but using a reagent that will not later be toxic to bacteria?
I think the best approach is irradiation, either UV or gamma irradiation or even X-ray, depending upon what you have access to. Heat would work with some phage but not necessarily with all. Some small phages survive at 95c although many are inactivated at 65C
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Hi all,
We are embarking on a journey into the exploration of the respiratory phageome. To test our methods we are collecting representative bacteriophage. One has eluded us so far and that is Acholeplasma virus L2 or a close relative. There does not appear to be a culture collection which includes this bacteriophage for purchase.
Is there are research group who would be willing to share an isolate and its propagation host of this elusive phage?
We would be very much grateful.
Thank you!
Chris
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I am looking to know the answer with best regards.
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Can anyone help me with protocol for removing host DNA contamination from phage lysate. Any protocol by using silica gel matrix for removal of host DNA
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use turbonuclease to remove host DNA.
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Lysate prep: donor strain is inoculated overnight in LB then diluted 1:100 into 5 mL LB + 0.2% glucose + 5 mM CaCl2, incubated for 1 hour. 100 uL of P1vir was added, then incubated until total lysis was seen. 200 uL of chloroform was added, vortexed, and spun down. Supernatant was kept and lysate was titered (~5E8 PFU/mL for most lysates).
Transduction: 5 mL overnight is inoculated in LB. Stationary culture is pelleted and resuspended in 2.5 mL of 10 mM MgCl2, 5 mM CaCl2. 100 uL cells was mixed with between 1-100 uL of P1 lysate. Phage was allowed to adsorb for 30 minutes at 37 C, after which 1 mL of LB + 200 uL of 1 M NaCitrate was added and incubated at 37 C with shaking for 1 hour. Cells were pelleted then plated on selective plates containing 5 mM NaCitrate as well as spotted onto nonselective plates containing 5 mM NaCitrate (controls). I'm also using a control with no phage added.
All solutions are autoclaved.
I originally got a few phage transductions to work (confirmed via PCR) however newer attempts have not succeeded. No colonies on any of my selective plates. On my nonselective LB plate, I notice a full growth in the no-phage-control spot, but only a couple of colonies in my added phage spots-- it seems that there is a lot of lysis going on. There does not seem to be a dose dependent response depending on how much phage I add (ie 1 uL of phage lysate added shows same result as 100 uL phage lysate added). I don't think I have phage contamination because I use same solutions and steps for the no-phage-control.
I've tried re-making solutions a few times with no avail either. Most confusingly, I retried using the lysate preps that were successful for me in the past and was unsuccessful this time around. There was also too much lysis as seen by spotting onto nonselective plates.
I'm thinking the only variables that there could be are the solutions, the phage lysate, the handling, and the recipient strain. It seems to me that there should not be variation in the phage lysates as I had tried lysates that were successful for me, and recipient strains are also the same. So that leaves handling/execution and solutions. I have remade solutions a few times now, and I have not changed the way I do transductions so I'm at a loss.
What is my next step in troubleshooting? Does anyone have insights into what could be going wrong?
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your strain background will not cause any problems. When I had this problem before I did the following:
1- take 3-5 lysates that worked previously and mix in a separate eppie all of them in equal volume then use that for preparing a new lysate.
2- Grow the lysate on a wild-type K12 strain, MG1655 should work.
3- Use the lysate from the MG1655 to make a new lysate from the keio strain
4- Repeat the transduction.
In case your out of luck with the above, you might want to consider getting a new P1 stock either from a neighboring lab or a culture collection.
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I need to get some phiX-174 and MS2 (the phages, not the DNA).
Can anyone suggest a supplier?
Andrew Jenkins, USN, Norway
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Thank you both for your help. Andrew
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Antimicrobial resistance is increasing despite new treatments being employed. With a decrease in the discovery rate of novel antibiotics, this threatens to take humankind back to a “pre-antibiotic era” of clinical care. Bacteriophages (phages) are one of the most promising alternatives to antibiotics for clinical use.
tell what is your thought on this and do you think we have a better solution than phage?
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The threat of antibiotic-resistant bacteria is pushing researchers to study other viable options. Bacteriophages, an old discovery widely disregarded after the discovery of antibiotics, are now being re-evaluated by ASU researchers. Phage therapy is the therapeutic use of bacteriophages as a treatment for bacterial infections. Each phage is different from the next. Researchers are currently trying to alter phages, and the next step would be to engineer phages. In order to fight bacteria, they must be engineered to fight different kinds of diseases for maximum effectiveness. At the moment, phage therapy is a mixture of different phages.
Also, kindly check the following link that may be useful:
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Tell me your ideas and thought on what you think is the best alternative to antibiotic sliver nanoparticle or bacteriophage.
in my opinion I think sliver nanoparticle is far better than phage because it is environmental friendly and it is cost effective.
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Nanotechnology continues to grow.Understanding the interaction of nanoparticles with bacteria and viruses is important to protect public health and enviroment .Effect of two commonly used nanoparticles Silver nanoparticles(AgNPs) average particles size 21nm on growth of bacteria and bacteriophage and applied for medicinal and enviromental uses.For detailed see the attached Ref.:
Hhttps://pub med.ncbi.gov.>
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I am analyzing somatic and MS2 coliphages in water using the single agar layer (SAL) method. The positive controls are determined using the double agar layer (DAL) method on a set of serial dilutions to find the dilution that will yield a target number of phages. It works perfectly for the MS2, but the somatic, even when the DAL is perfect, yields 500-600% when I run the SAL along with my samples. Calculations are correct, there is no contamination, I tried filtering the somatic dilution through a 22 micron sterile syringe filter before adding (to decrease the possibility of phage clumping in the dilution). I still get >500% recovery.
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If you use the exact same dilution in both your SAL and DAL counts, do you get the same value or not. If not, then it clearly is a procedural problem either by not visualizing many plaques (perhaps small ones) or differences in efficiency in phage absorption with each type.
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Hello everyone.
I am a beginner in phage display of antibodies. We have an in-home prepared phage display antibody library. I performed biopanning against an antigen, a purified protein, with phage expressing the library. In order to make the selection stringent, I performed 60x washings 1minute each (30x with PBS-T 0.1% tween and 30x with PBS).
After biopanning, when comparing to number of eluted phages in Antigen coated vs No Antigen wells, I got ~70 fold more colonies in Antigen coated wells compared to No Antigen well.
{Short Summary}
When I performed monoclonal phage ELISA, I could not observe any difference between No-Ag vs Ag well for each clone. Infact, in some cases the OD was higher in No-Ag as compared to Ag. Seeing that the enrichment at biopanning level was nice, it is really disheartening to see none of the clones reacted. What could be the possible reasons ? Could it be that the antibodies might cross react with blocking solution components ? Or is it that none of the clone binds ?
{Detailed Procedure}
Therefore, I proceeded with colonies that have correct antibody fragment (based on colony PCR with vector specific primers) and performed a monoclonal phage ELISA. For monoclonal phage ELISA, 1mL secondary culture was set up from primary culture of each colony. The secondary culture was then infected with helper phages OD600=0.5 at 1:20 infection ratio and then Kan(helper phage) and IPTG were added. This culture was then grown @ 37degC under shaking for 9-10 hours. Phages were then purified with PEG/NaCl and then used for phage ELISA.
For each clone, 3wells were No-Ag while 3-wells were Ag coated. Purified phages were diluted in blocking solution and divided equally in all 6-wells, following blocking the wells. Phages were detected with anti-M13KO7 Hyper Immune Serum and further detected with anti-Mouse AP conjugated antibody.
I could not observe any difference between No-Ag vs Ag well for each clone. Infact, in some cases the OD was higher in No-Ag as compared to Ag.
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Abhishek Dubey I see, that should work fine. You did it in triplicate, is it replicable? That is, when you have higher signal for no-Ab than for Ab is it the same for all three wells?
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Hello, research community.
Several genes are found to be present in bacteria and bacteriophages. For example, the FlgB gene is present in Campylobacter phage and their host Campylobacter jejuni as well as in E. coli. Then, the stxA, stxB genes are found in E. coli bacteria and their predator bacteriophages. Going through several articles, I have found all the three genes mentioned above are of bacterial origin and transferred to bacteriophages.
Now, I would like to know if there is any tool or method available to identify if the gene is evolved in bacteria and transferred to bacteriophages or vice versa.
Thank you.
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Collect all the homologues of the gene from all available organisms including the bacterial and viral strains and perform a multiple sequence alignment. Then build a phylogenetic tree and analyze the distance from your target organism to get an idea where the gene evolved from. Clustal Omega and PROBCONS are some tools for basic alignment. They also provide an option to visualize the tree. MEGA tree explorer and Phylotree are some tools for analyzing the tree.
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The plate is made with 2TY with Ampicillin, Kanamycin and Glucose. The plaques that form are small and big on the same petri dish, its got a pretty disproportionate look to it.
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Michael J. Benedik The issue is actually different sizes on a single plate, but thanks for your help!
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Hello I am currently trying to amplify helper phage VCSM13 with TG1 E.coli.
The problem is that I have tried very hard to amplify several times but it all failed, ending with no plaques. (LB agar + Top agar)
So I have tried to take phage in supernatant in every step of the protocol, and plated in LK plate.
The colonies were seen fine, even until the final step.
However, when I plate in on top agar for titering, there are no plaques seen.
I can see that infection works okay (by LK plate), but will there be any reason why there are no plaque formation?
Also, at the first day of the protocol to get take a single plaque from the stock which we want to amplify, the plaques form well after overnight.
So to put it short, the plaques form well on day 1 directly from the stock, but after amplification, no plaque is formed.
(I can titer with CFU, but not PFU)
Would it just be the problem of infection efficiency of the phage stock?
It would be very helpful if there would be any advice. Thank you.
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Hi Hye nice question.
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When I spot the phage lysate on MG1655, I see the clearance. But, to calculate the phage titer I mix the lysate (100ul) with MG1655 as background strain (100ul) in 3ml 0.5% agar. In this case, I am not getting any plaques. What could be the possible reasons?
I diluted my overnight culture of the background strain to OD 0.1 and then used this in the top agar method.
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Because absence of microorgnism or lack or defect in stain crystal violet or inubation temperatue.
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A probably silly question to phage experts.
Is it possible to determine whether a phage genome is linear, circular, or circularly permutated using Illumina sequencing data and genome assembly? Thank you very much!
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Greetings,
Elucidation of the exact genome termini is what you probably want.
I've found PhageTerm (Garneau et al., 2017) to be a great tool to have an automated look at the phage read pile-ups onto the scaffold representing phage complete genome for termini determination, which works well with paired reads when the library prep involved random physical shearing (we usually use TruSeq DNA Nano workflow with shearing by Covaris ultrasonication; however, it won't be suitable for Nextera-prepared libraries, which is a very common library prep choice). Even of case of lackluster coverage, read pile-up inspection and TerL aa sequence tree generation with your phage TerL sequence and other TerL sequences from phages that had their termini elucidated experimentally (not only in silico) can usually aid in the proper in vitro experiment design (have a look at Casjens et al., 2009) to verify/find out the packaging strategy/termini phage has.
Best regards,
Nikita Zrelovs
1. Garneau, J.R., Depardieu, F., Fortier, LC. et al. PhageTerm: a tool for fast and accurate determination of phage termini and packaging mechanism using next-generation sequencing data. Sci Rep 7, 8292 (2017). https://doi.org/10.1038/s41598-017-07910-5
2. Casjens, Sherwood R, and Eddie B Gilcrease. “Determining DNA packaging strategy by analysis of the termini of the chromosomes in tailed-bacteriophage virions.” Methods in molecular biology (Clifton, N.J.) vol. 502 (2009): 91-111. doi:10.1007/978-1-60327-565-1_7
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