SEA-PHAGES | All posts created by viknesh

Link to this post \| posted 12 Nov, 2021 14:54
viknesh	afreise Hi everyone, Also, I'd be interested to hear if anyone at SEA-PHAGES HQ has priority hosts they'd like new phages on. Thanks in advance for any thoughts! Amanda Hi Amanda, Reiterating what Debbie said that 1. we would be supportive of anyone exploring new Actino hosts 2. we have limited experience with hosts that we've not already shared with the SEA For the recent lot of new hosts that are being used in the program, we spent a year with each before before introducing it to everyone. We primarily looked for two things… 1) can we grow it on media we already have (7H9 or PYCa) with minimal modifications, and 2) can we isoalte phage at at least 10 % of samples tested. When a host can meet these criteria, we introduce it into the program because it means faculty can wrk with it and that students can have some success with it. When it doesnt meet these criteria, we dont share news about it. For example,Corynebacterium flavescens is one that was easy to grow but SO hard to find phage on (1 in 50 samples yileded phage). We used to have this information on the website… perhaps it is time to update it! If you are willing to work with new hosts, I am certainly happy to have a discussion with you about which you could try (and even send you those strains, if we have them). However, I would recommend that this happen outside of the PHAGES project until we know it is something students can have some success with. Happy to chat. Vic

Link to this post | posted 12 Nov, 2021 14:54

afreise
Hi everyone,

Also, I'd be interested to hear if anyone at SEA-PHAGES HQ has priority hosts they'd like new phages on.

Thanks in advance for any thoughts!
Amanda

Hi Amanda,

Reiterating what Debbie said that
1. we would be supportive of anyone exploring new Actino hosts
2. we have limited experience with hosts that we've not already shared with the SEA

For the recent lot of new hosts that are being used in the program, we spent a year with each before before introducing it to everyone. We primarily looked for two things…
1) can we grow it on media we already have (7H9 or PYCa) with minimal modifications, and
2) can we isoalte phage at at least 10 % of samples tested.
When a host can meet these criteria, we introduce it into the program because it means faculty can wrk with it and that students can have some success with it. When it doesnt meet these criteria, we dont share news about it. For example,Corynebacterium flavescens is one that was easy to grow but SO hard to find phage on (1 in 50 samples yileded phage). We used to have this information on the website… perhaps it is time to update it!

If you are willing to work with new hosts, I am certainly happy to have a discussion with you about which you could try (and even send you those strains, if we have them). However, I would recommend that this happen outside of the PHAGES project until we know it is something students can have some success with.

Happy to chat.
Vic

Posted in: Phage Discovery/Isolation → New hosts - human microbiome

Link to this post \| posted 27 Oct, 2021 21:43
viknesh	stpage Hi, the phage discovery has RNase/DNase/ProteinaseK treatment followed by CleanupResin (guanidinium thiocyanate) to denature the capsid. Some protocols seem to use proteinaseK to denature the capsid. (https://www.mdpi.com/2409-9279/1/3/27/pdf-vor) and some seem intermediate (https://pubmed.ncbi.nlm.nih.gov/29417429/). Is it very phage dependent? With new hosts do we need to optimize the proteinaseK treatment?? Hi Shallee, The guanidinium thiocyanate in the cleanup resin is enough to denature phage capsids, at least for all the phages we work with. If you or anyone else is having trouble with DNA isolation, please let us know so we can look into this. As for the additional "optional" ProteinaseK + SDS step in the guide, this was added because we were observing some nuclease activity co-purifying with DNA for some phages isolated on M. foliorum. You'll know if you have this problem if your phage DNA looks good on a gel if the sample hasnt seen any cations (which are needed for DNase activity), but is degraded if it has. For example, even the addition of restricton enzyme buffer (i.e. for the DNA control with no actual restriction enzyme) would result in degradation of phage DNA, but loading the DNA sample directly onto the gel results in the typical nice high-molecular-weight band (with little to no smear). While we dont know why nuclease co-purifies for M. foliorum phages, or even what the nuclease is (whether its the nuclease we add or some M. foliorum nuclease), adding ProteinaseK and SDS after treating lysates with nuclease resolves the problem. As you'll recall, we typically simply inactivate the nucleases by adding EDTA. For those working with M. foliorum, or for those seeing nuclease activity co-purifying with their phage DNA, the addition of Proteinase K and SDS that is able to degrade that nuclease can resolve the problem. I hope this is helpful. Vic

Link to this post | posted 27 Oct, 2021 21:43

viknesh

stpage
Hi, the phage discovery has RNase/DNase/ProteinaseK treatment followed by CleanupResin (guanidinium thiocyanate) to denature the capsid.
Some protocols seem to use proteinaseK to denature the capsid. (https://www.mdpi.com/2409-9279/1/3/27/pdf-vor)
and some seem intermediate (https://pubmed.ncbi.nlm.nih.gov/29417429/).

Is it very phage dependent? With new hosts do we need to optimize the proteinaseK treatment??

Hi Shallee,

The guanidinium thiocyanate in the cleanup resin is enough to denature phage capsids, at least for all the phages we work with. If you or anyone else is having trouble with DNA isolation, please let us know so we can look into this.

As for the additional "optional" ProteinaseK + SDS step in the guide, this was added because we were observing some nuclease activity co-purifying with DNA for some phages isolated on M. foliorum.
You'll know if you have this problem if your phage DNA looks good on a gel if the sample hasnt seen any cations (which are needed for DNase activity), but is degraded if it has. For example, even the addition of restricton enzyme buffer (i.e. for the DNA control with no actual restriction enzyme) would result in degradation of phage DNA, but loading the DNA sample directly onto the gel results in the typical nice high-molecular-weight band (with little to no smear).

While we dont know why nuclease co-purifies for M. foliorum phages, or even what the nuclease is (whether its the nuclease we add or some M. foliorum nuclease), adding ProteinaseK and SDS after treating lysates with nuclease resolves the problem. As you'll recall, we typically simply inactivate the nucleases by adding EDTA. For those working with M. foliorum, or for those seeing nuclease activity co-purifying with their phage DNA, the addition of Proteinase K and SDS that is able to degrade that nuclease can resolve the problem.

I hope this is helpful.

Vic

Posted in: Phage Discovery/Isolation → Capsid denaturation

Link to this post \| posted 27 Sep, 2021 14:56
viknesh	cheryl.brown@mnstate.edu Thank you. As a follow-up, are you aware of anyone else having difficulties with VWR filters recently? We have been using the same filters for 3 years and this fall is the first time we have had the problem. It seems like a QA failure. We ran a test with the current filters and some others on hand, and, using the same force in both, the other filters (also .22 micron syringe filters), only the VWR ones had the contamination problem. I have some .2 micron PES filters on hand. Can those be used in this application? We are also going to tweak the pre-filter/coffee filter step to further decrease particles in samples. I've personally not heard any recent complaints about VWR filters, but perhaps others will chime in. The 0.2 um PES filters will work fine. Vic

Link to this post | posted 27 Sep, 2021 14:56

viknesh

cheryl.brown@mnstate.edu
Thank you. As a follow-up, are you aware of anyone else having difficulties with VWR filters recently? We have been using the same filters for 3 years and this fall is the first time we have had the problem. It seems like a QA failure.

We ran a test with the current filters and some others on hand, and, using the same force in both, the other filters (also .22 micron syringe filters), only the VWR ones had the contamination problem.

I have some .2 micron PES filters on hand. Can those be used in this application?

We are also going to tweak the pre-filter/coffee filter step to further decrease particles in samples.

I've personally not heard any recent complaints about VWR filters, but perhaps others will chime in. The 0.2 um PES filters will work fine.

Vic

Posted in: Phage Discovery/Isolation → Syringe filter failure?

Link to this post \| posted 26 Sep, 2021 19:51
viknesh	cheryl.brown@mnstate.edu We have been plagued with recurrent contamination problems on plates. One of the confusing factors was the fact that the populations of contaminants was varied. We have widespread, but inconsistent, contamination. I just ran a test yesterday, following the student protocols. I soaked the soil, ran it through a coffee filter, and then through a syringe filter. One sample was more plant material, so the broth needed no force to go through. The other sample was a fine, silty soil, and considerable pressure was needed to get it filtered. There was no obvious break or breach, as the same pressure was needed the entire time. I then plated the filtrate with sterile top agar and no culture. The "forced" plate was covered with bacteria. Is there a change in protocol needed? Additional pre-filtering? Hi Cheryl, These filters are not designed to work under pressure. As soon as it becomes difficult to push liquid through the filter, the filter pores are essentially clogged/blocked and you've reached the filtration capacity of that filter. Even with steady force, you risk micro-ruptures and eventually a full rupture of the membrane. For clay or silty samples, which always clog the filters, I spin the soil suspension (in 50 ml conical tubes) at 4 krpm 10 minutes. This is usually fast enough to pellet much of the particulate matter so that I can avoid clogging the filters too quickly. Once it clogs, I either use a new filter or just enrich with however little filtrate I am able to get. Often, it is easy to obtain at least 10 ml of filtrate before these samples (spun at 4krpm) clog the filter. I hope this is helpful. Vic

Link to this post | posted 26 Sep, 2021 19:51

viknesh

cheryl.brown@mnstate.edu
We have been plagued with recurrent contamination problems on plates. One of the confusing factors was the fact that the populations of contaminants was varied. We have widespread, but inconsistent, contamination.

I just ran a test yesterday, following the student protocols. I soaked the soil, ran it through a coffee filter, and then through a syringe filter. One sample was more plant material, so the broth needed no force to go through. The other sample was a fine, silty soil, and considerable pressure was needed to get it filtered. There was no obvious break or breach, as the same pressure was needed the entire time. I then plated the filtrate with sterile top agar and no culture.

The "forced" plate was covered with bacteria.

Is there a change in protocol needed? Additional pre-filtering?

Hi Cheryl,

These filters are not designed to work under pressure. As soon as it becomes difficult to push liquid through the filter, the filter pores are essentially clogged/blocked and you've reached the filtration capacity of that filter. Even with steady force, you risk micro-ruptures and eventually a full rupture of the membrane.

For clay or silty samples, which always clog the filters, I spin the soil suspension (in 50 ml conical tubes) at 4 krpm 10 minutes. This is usually fast enough to pellet much of the particulate matter so that I can avoid clogging the filters too quickly. Once it clogs, I either use a new filter or just enrich with however little filtrate I am able to get. Often, it is easy to obtain at least 10 ml of filtrate before these samples (spun at 4krpm) clog the filter.

I hope this is helpful.

Vic

Posted in: Phage Discovery/Isolation → Syringe filter failure?

Link to this post \| posted 20 Sep, 2021 14:44
viknesh	cheryl.brown@mnstate.edu Hello all, Recently we have been having more issues with bubbles in our agar plates. I have extended the "rest" time after adding CaCl2 and dextrose to allow any foam from swirling to dissipate, but still getting bubbles in my plates. Is there an additive that works with PYCa and M. foliorum that is anti-foam or anti-bubble? Do any of you use it or have you? Hi Cheryl, It sounds like these bubbles form when you swirl to mix calcium and dextrose into the molten agar. If this is true, then my recommendation is to swirl to mix gently. We typically have a stir bar in the bottle (added before we autovlave the media) that is set to spin slow enough to not create any bubbles. If you are swirling by hand, simply rotating or rolling the bottle on its side should be sufficient mixing. Because the media is so viscous, especially when its not piping hot, it is hard to get rid of any bubbles that form without having to reheat the media. I hope this is helpful. Vic Edited 20 Sep, 2021 14:46

Link to this post | posted 20 Sep, 2021 14:44

viknesh

cheryl.brown@mnstate.edu
Hello all,

Recently we have been having more issues with bubbles in our agar plates. I have extended the "rest" time after adding CaCl2 and dextrose to allow any foam from swirling to dissipate, but still getting bubbles in my plates.

Is there an additive that works with PYCa and M. foliorum that is anti-foam or anti-bubble? Do any of you use it or have you?

Hi Cheryl,

It sounds like these bubbles form when you swirl to mix calcium and dextrose into the molten agar. If this is true, then my recommendation is to swirl to mix gently. We typically have a stir bar in the bottle (added before we autovlave the media) that is set to spin slow enough to not create any bubbles. If you are swirling by hand, simply rotating or rolling the bottle on its side should be sufficient mixing. Because the media is so viscous, especially when its not piping hot, it is hard to get rid of any bubbles that form without having to reheat the media.

I hope this is helpful.

Vic

Edited 20 Sep, 2021 14:46

Posted in: Phage Discovery/Isolation → Anti-foam or anti-bubble additives?

Link to this post \| posted 10 Sep, 2021 17:55
viknesh	c.sunnen So after running this course for 6+ years, I discovered that my colleague and I clean-up our benches differently at the end of lab, and we both swear we were taught (during the same training) the way we each do it is "correct." Whenever disinfecting BEFORE lab, we always do Ci-decon first, let it dry, followed by 70% Ethanol. The question is whether the order should be reversed at the end of experiments, before leaving the lab. Should it remain Ci-decon first, then EtOH? Or should it be EtOH first, followed by Ci-decon? Does it matter? (and if it does matter, can you share why?) We typically do EtOH after CiDecon so that we remove from the bench residual phenol from the cidecon.

Link to this post | posted 10 Sep, 2021 17:55

viknesh

c.sunnen
So after running this course for 6+ years, I discovered that my colleague and I clean-up our benches differently at the end of lab, and we both swear we were taught (during the same training) the way we each do it is "correct."

Whenever disinfecting BEFORE lab, we always do Ci-decon first, let it dry, followed by 70% Ethanol. The question is whether the order should be reversed at the end of experiments, before leaving the lab.

Should it remain Ci-decon first, then EtOH? Or should it be EtOH first, followed by Ci-decon? Does it matter?

(and if it does matter, can you share why?)

We typically do EtOH after CiDecon so that we remove from the bench residual phenol from the cidecon.

Posted in: Phage Discovery/Isolation → Disinfection Debate: does the order matter?

Link to this post \| posted 14 Apr, 2021 19:25
viknesh	kmaclea I wanted to send an update that we have conclusively proven the source of our contamination giving us so many problems. The agar we were using to make the top agar appears to have been contaminated with something that could survive 40 minutes of autoclaving (volume 500 ml). Now that we have switched out the agar, our contamination issues have gone away completely. Just an update for anyone following the thread. Kyle Thrilled to hear it, Kyle. Hope your students had fun figuring this out.

Link to this post | posted 14 Apr, 2021 19:25

viknesh

kmaclea
I wanted to send an update that we have conclusively proven the source of our contamination giving us so many problems.

The agar we were using to make the top agar appears to have been contaminated with something that could survive 40 minutes of autoclaving (volume 500 ml).

Now that we have switched out the agar, our contamination issues have gone away completely.

Just an update for anyone following the thread.

Kyle

Thrilled to hear it, Kyle. Hope your students had fun figuring this out.

Posted in: Phage Discovery/Isolation → Contamination of top agar and possibly other components

Link to this post \| posted 03 Apr, 2021 16:57
viknesh	kmaclea So here's an update. https://docs.google.com/document/d/1294ZwYilXMbQA6n0eFrWouodmvBk1M4ZOL4AsPiUsX8/ Thanks for sharing all this, Kyle. I can't say for sure, of course, but it looks like you might have multiple things going on. First - the media. Did you try imaging the most recent cloudy bottle of top agar (the one that your TA added calcium and dextrose before authoclaving?). I ask because CaCl2 precipitates when heated for prolonged preiods (i.e. autoclaving), and the amount of precipitation depends on the pH and other salts in solution. While I am not discounting your contamination (all those beautiful microscopy images), the cloudiness in the recent bottles might just be precipitate, and cloudy top agar may be sufficient for plaques to look cloudy - in which case your contamination problems might be isolated events and not a widespread problem. Second - I think those contaminated spots on the enriched plates are not coming from the top agar, but instead from the samples spotted themselves. This is fairly common when the enrichment is not properly filtered, typically because there has been a rupture in the filter membrane. I used to see this fairly often and so now buy the fairly "tough" GDX Whatmann filters that rupture less often (but which still occasionally do). Of course, maybe you have mycoplasma, which can pas through the filter). For your plaque assay plates that are very clearly contaminated, again, this might be a problem with the sample itself being contaminated since some other plates look great - are these also filtered soil samples? If you performed/perform a dilution series with these samples, my guess if that you'll see the contamination dilute out too. Kyle, regardless of what's happening, the fact hat you're engaging your students in trying to figure out this issue is pretty awesome! What a ride for them Vic Edited 03 Apr, 2021 16:59

Link to this post | posted 03 Apr, 2021 16:57

viknesh

kmaclea
So here's an update.
https://docs.google.com/document/d/1294ZwYilXMbQA6n0eFrWouodmvBk1M4ZOL4AsPiUsX8/

Thanks for sharing all this, Kyle. I can't say for sure, of course, but it looks like you might have multiple things going on.

First - the media. Did you try imaging the most recent cloudy bottle of top agar (the one that your TA added calcium and dextrose before authoclaving?). I ask because CaCl2 precipitates when heated for prolonged preiods (i.e. autoclaving), and the amount of precipitation depends on the pH and other salts in solution. While I am not discounting your contamination (all those beautiful microscopy images), the cloudiness in the recent bottles might just be precipitate, and cloudy top agar may be sufficient for plaques to look cloudy - in which case your contamination problems might be isolated events and not a widespread problem.

Second - I think those contaminated spots on the enriched plates are not coming from the top agar, but instead from the samples spotted themselves. This is fairly common when the enrichment is not properly filtered, typically because there has been a rupture in the filter membrane. I used to see this fairly often and so now buy the fairly "tough" GDX Whatmann filters that rupture less often (but which still occasionally do). Of course, maybe you have mycoplasma, which can pas through the filter).

For your plaque assay plates that are very clearly contaminated, again, this might be a problem with the sample itself being contaminated since some other plates look great - are these also filtered soil samples? If you performed/perform a dilution series with these samples, my guess if that you'll see the contamination dilute out too.

Kyle, regardless of what's happening, the fact hat you're engaging your students in trying to figure out this issue is pretty awesome! What a ride for them smile

Vic

Edited 03 Apr, 2021 16:59

Posted in: Phage Discovery/Isolation → Contamination of top agar and possibly other components

Link to this post \| posted 31 Mar, 2021 11:52
viknesh	kmaclea So, this is going to sound insane, but we've been running into recurrent bacterial contamination that a couple of attempts to fix doesn't seem to be fixing. We are trying to use both A. globiformis and M. foliorum for phage discovery. We had a lot of luck getting plaques originally, then at some point most of our students' phages have been unable to propagate further (no plaques). We've done countless direct and enriched isolations. A few weeks ago we had a contamination issue and re-made components. Things got better for a time, but then we were back in problems again. Last week there was obvious rampant contamination again. We received new glycerol stocks for bacteria, remade all the PYCa, all the top agar, all the dextrose, CaCl2, and CHX. The last three were sterile filtered, the first two were autoclaved. Autoclaving was standardly done with 500 ml at a time and 45 minute autoclaving at 121C. By all indications, other media being made using the same autoclave is just fine although no biological testing of the autoclave has been done recently. Most recent tests seem to indicate it is the top agar in which we are seeing contamination. Even in freshly made top agar, freshly autoclaved, with the other components all sterile filtered in the hood. Attention being paid to sterile technique during the handling, in the hood. What could be making our top agar contaminated? We have been using a metal bead bath for incubation of the top agar, but we are seeing this even in freshly made top agar. Beads in the bead bath have also been recently decontaminated. Ideas of any kind would be most definitely welcomed! Kyle Hi Kyle, Looks like there ate at least two problems: 1) not being abe to propoagate a phage and 2) contaminated materials. For #1, are you seeing this for both hosts, or just one? If the latter, which one. Also, will the control phage also not plaque? For #2, are you able to share images of the contamination? I can;t tell if the top agar is getting contaminated as you use it or whether it was never actually fully sterilized. For example, what happens to the freshly autoclaved top agar, still molten in the bottle with the lid closed, if left in the beadbath for a few days - never opened? Do you see microbial growth in the bottle, or only when you use it? Vic

Link to this post | posted 31 Mar, 2021 11:52

viknesh

kmaclea
So, this is going to sound insane, but we've been running into recurrent bacterial contamination that a couple of attempts to fix doesn't seem to be fixing.

We are trying to use both A. globiformis and M. foliorum for phage discovery. We had a lot of luck getting plaques originally, then at some point most of our students' phages have been unable to propagate further (no plaques). We've done countless direct and enriched isolations.

A few weeks ago we had a contamination issue and re-made components. Things got better for a time, but then we were back in problems again. Last week there was obvious rampant contamination again. We received new glycerol stocks for bacteria, remade all the PYCa, all the top agar, all the dextrose, CaCl2, and CHX. The last three were sterile filtered, the first two were autoclaved. Autoclaving was standardly done with 500 ml at a time and 45 minute autoclaving at 121C. By all indications, other media being made using the same autoclave is just fine although no biological testing of the autoclave has been done recently.

Most recent tests seem to indicate it is the top agar in which we are seeing contamination. Even in freshly made top agar, freshly autoclaved, with the other components all sterile filtered in the hood. Attention being paid to sterile technique during the handling, in the hood.

What could be making our top agar contaminated? We have been using a metal bead bath for incubation of the top agar, but we are seeing this even in freshly made top agar. Beads in the bead bath have also been recently decontaminated.

Ideas of any kind would be most definitely welcomed!

Kyle

Hi Kyle,

Looks like there ate at least two problems:
1) not being abe to propoagate a phage and
2) contaminated materials.

For #1, are you seeing this for both hosts, or just one? If the latter, which one. Also, will the control phage also not plaque?
For #2, are you able to share images of the contamination? I can;t tell if the top agar is getting contaminated as you use it or whether it was never actually fully sterilized. For example, what happens to the freshly autoclaved top agar, still molten in the bottle with the lid closed, if left in the beadbath for a few days - never opened? Do you see microbial growth in the bottle, or only when you use it?

Vic

Posted in: Phage Discovery/Isolation → Contamination of top agar and possibly other components

Link to this post \| posted 08 Feb, 2021 01:21
viknesh	kmaclea Dear Colleagues We are running phage discovery in the winter for the first time. Our first go-round we have a lot of success with direct isolations out of the box. This might have been an outlier leading us astray, but over the last couple of weeks we only had one successful run with direct isolation. We have started enriched isolation but still wondering if we can do better with our direct isolation or if winter in New Hampshire is just going to be more challenging. Anyone have thoughts? Kyle Hi Kyle, From what I can tell based on findings from across the SEA, the success rate for direct isolation is generally low and inconsistent (unless you keep going back to the same/similar spot). I'll let faculty from the northern reaches of the country share their advice, but I know several who gather soil early (before the ground freezes, and store them in sealed ziplocks at 4C) or use soil from indoor pots. I personally have had success with both methods when isolating phage for M. foliorum, but only by enriched isolation. Good luck!

Link to this post | posted 08 Feb, 2021 01:21

viknesh

kmaclea
Dear Colleagues

We are running phage discovery in the winter for the first time. Our first go-round we have a lot of success with direct isolations out of the box. This might have been an outlier leading us astray, but over the last couple of weeks we only had one successful run with direct isolation. We have started enriched isolation but still wondering if we can do better with our direct isolation or if winter in New Hampshire is just going to be more challenging.

Anyone have thoughts?

Kyle

Hi Kyle,

From what I can tell based on findings from across the SEA, the success rate for direct isolation is generally low and inconsistent (unless you keep going back to the same/similar spot). I'll let faculty from the northern reaches of the country share their advice, but I know several who gather soil early (before the ground freezes, and store them in sealed ziplocks at 4C) or use soil from indoor pots. I personally have had success with both methods when isolating phage for M. foliorum, but only by enriched isolation.

Good luck!

Posted in: Phage Discovery/Isolation → Phage isolation in winter--any differences?

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