Bacteriophage - Science topic

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!

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.

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

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.

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).

Dear Michael J. Benedik , thank you for your valuable response and advice.

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.

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.

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

Michael J. Benedik and Pierre Béguin thank you for your answers. I will proceed with my experiments keeping the given suggestions in mind.

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.

I agree with Dr. Michael J.Benedik answer

how can I create drought conditions by using PEG 6000 for drought-resistant rhizobacteria in the laboratory to check their drought-resistant attributes

Malavika B H Wow, yes 10¹³ is a very high titre. That's not the problem then. Not sure what else to suggest.

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

You may also try reducing the percentage of top agar

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.

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.

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.

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,

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.

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:

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:

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.

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!

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.

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.

Also, kindly check:

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

DOI:https://doi.org/10.1128/JB.01080-13ee this link

Best Regard.

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.

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

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:

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.

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".

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.

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).

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.

I am so grateful for the data. it has been of great use

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.

Hi James Bruns .

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!

Your top agar is not well dissolved.

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.

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.

Bacterial growth occur on phage plate due to strain specifity and O.D culture.

Hi Hyago Passe Pereira ,

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!

That makes sense. Thank you very much.

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.

Thank you!

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.

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

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.

iTRAQ-based Proteomics Analysis may help you at initial level.

Thankyou

https://academic.oup.com/femsmicrobes/article/doi/10.1093/femsmc/xtab003/6204716#235345021

Hope this paper will help you for isolation. Isolation and characterization of bacteriophages from the human skin microbiome that infect Staphylococcus epidermidis.

I would like to know the detailed protocol, especially about Cas9-gRNA preparation.

Michael J. Benedik unfortunately no picture with me now, but I will post it definitly in my upcoming manipulations. thanks for your recommendation.

Macrogene

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

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.

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.

Thank you will do!!!

thanks wolfgang, i've been considering probiotic strains previously

I think the identification of regulatory elements, primarily promoters, which are specific DNA regions responsible for transcription initiation. Muskan Gupta

Thanks. let me know if you learn anything new on phi6 propagation, Im still struggling with it.

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

I am looking to know the answer with best regards.

use turbonuclease to remove host DNA.

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.

Thank you both for your help. Andrew

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:

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.>

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.

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?

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.

Michael J. Benedik The issue is actually different sizes on a single plate, but thanks for your help!

Hi Hye nice question.

Because absence of microorgnism or lack or defect in stain crystal violet or inubation temperatue.

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