SEA-PHAGES | All posts created by viknesh

Link to this post \| posted 13 Sep, 2022 23:38
viknesh	cellokiwi So when you pass it, it goes back to yellow for a bit then turns orange again. We sadly don't have the resources to check 16S. Is there a resource for this through the program? The first time we saw it was in a lysogen so I just attributed it to wherever the genome had integrated itself messing with the color. Now, there has been zero change whatsoever. Same media, same CaCl2, same buffer, same incubator, same everything. One colleague suggested it is a carotenoid response to oxidative stress? Does that sound possible? Since we're using the same everything what would suddenly be stressing them out that wasn't before? In the picture the one on the left is a normal Arthrobacter culture and the one on the right is just starting to turn orange. They get darker than that. Hi Alison, Since we began using M . foliorum for phage-hunting in the SEA, I've yet to hear from anyone about it turning orange. Can you confirm the following? 1. If you streak a plate from your freezer/glycerol stock and incubate the plate at 30C, colonies start out yellow but then turn orange? How long before they turn yellow, and how long before they turn orange? If you have photos, please share. 2. If you setup a culture, they saturated culture is yellow but then eventually turns orange? If so, as before, please share the timing of these color changes. 3. If you streak a plate from the culture (from #2), colony formation and timin gof color changes proceed similar to streaking from the glycerol? 4. You have several phages that can still form plaques when plated with bacteria from the orange culture? Thanks. Vic Edited 13 Sep, 2022 23:39

Link to this post | posted 13 Sep, 2022 23:38

cellokiwi
So when you pass it, it goes back to yellow for a bit then turns orange again. We sadly don't have the resources to check 16S. Is there a resource for this through the program? The first time we saw it was in a lysogen so I just attributed it to wherever the genome had integrated itself messing with the color. Now, there has been zero change whatsoever. Same media, same CaCl2, same buffer, same incubator, same everything. One colleague suggested it is a carotenoid response to oxidative stress? Does that sound possible? Since we're using the same everything what would suddenly be stressing them out that wasn't before?

In the picture the one on the left is a normal Arthrobacter culture and the one on the right is just starting to turn orange. They get darker than that.

Hi Alison,

Since we began using M . foliorum for phage-hunting in the SEA, I've yet to hear from anyone about it turning orange. Can you confirm the following?

1. If you streak a plate from your freezer/glycerol stock and incubate the plate at 30C, colonies start out yellow but then turn orange? How long before they turn yellow, and how long before they turn orange? If you have photos, please share.

2. If you setup a culture, they saturated culture is yellow but then eventually turns orange? If so, as before, please share the timing of these color changes.

3. If you streak a plate from the culture (from #2), colony formation and timin gof color changes proceed similar to streaking from the glycerol?

4. You have several phages that can still form plaques when plated with bacteria from the orange culture?

Thanks.
Vic

Edited 13 Sep, 2022 23:39

Posted in: Arthrobacter → Orange Cells

Link to this post \| posted 23 Aug, 2022 16:20
viknesh	A for effort and yAy for a cluster that with congruent calls! debbie Hi all, As of today, June 17, 2022, there are 88 Cluster EE genomes in our records. They are closely related genomes that contain only ~28 genes. In an effort to make the records congruent, my students and I have reviewed ~75 of them and revised 71 of the records. The template we used is attached here, along with a list of 71 records that we touched. We spent an abundant amount of time on the helix-turn-helix DNA binding proteins at the right end of the genome. We investigated 'types' of helix-turn-helix choices and decided the best path is to call them helix-turn-helix DNA binding proteins. August 23, 2022 - I just received word that the genomes that we sent to GenBank have been processed. This cluster just might have congruent calls! Thanks Vic! debbie Edited 23 Aug, 2022 18:40

Link to this post | posted 23 Aug, 2022 16:20

viknesh

A for effort and yAy for a cluster that with congruent calls!

debbie
Hi all,
As of today, June 17, 2022, there are 88 Cluster EE genomes in our records. They are closely related genomes that contain only ~28 genes. In an effort to make the records congruent, my students and I have reviewed ~75 of them and revised 71 of the records. The template we used is attached here, along with a list of 71 records that we touched. We spent an abundant amount of time on the helix-turn-helix DNA binding proteins at the right end of the genome. We investigated 'types' of helix-turn-helix choices and decided the best path is to call them helix-turn-helix DNA binding proteins.
August 23, 2022 - I just received word that the genomes that we sent to GenBank have been processed. This cluster just might have congruent calls!

Thanks Vic!
debbie

Edited 23 Aug, 2022 18:40

Posted in: Cluster EE Annotation Tips → Genome Curation - a must read!

Link to this post \| posted 05 May, 2022 17:50
viknesh	bgibb There used to be three protocols for setting up TEM samples, but now there is only one 8.1c (parafilm drop method) in the Discovery guide and Instructor guide. Is there a reason that the other protocols were removed? I've had good luck with the pelco tab method. Bryan, I see all 3 protocols in the Discovery Guide - https://seaphagesphagediscoveryguide.helpdocsonline.com/8-0-toc

Posted in: Phage Discovery/Isolation → Electron microscopy protocols

Link to this post \| posted 25 Jan, 2022 15:50
viknesh	sahas Hi I was wondering if anyone can recommend a place/lab to send our samples for electron microscopy? We haven't been successful in finding a resource yet. Any help will be highly appreciated. Thanks Sangha Similar to Cathy's suggestion, you could also reachout to UMBC's Imaging Facility. If you would like to cehck on their pricing and availability, you can email their facility director, Tagide deCarvalho at tagided@umbc.edu. Please copy me on that email to Tagide.

Link to this post | posted 25 Jan, 2022 15:50

viknesh

sahas
Hi
I was wondering if anyone can recommend a place/lab to send our samples for electron microscopy? We haven't been successful in finding a resource yet.
Any help will be highly appreciated.
Thanks
Sangha

Similar to Cathy's suggestion, you could also reachout to UMBC's Imaging Facility. If you would like to cehck on their pricing and availability, you can email their facility director, Tagide deCarvalho at tagided@umbc.edu. Please copy me on that email to Tagide.

Posted in: General Message Board → Electron Microscopy

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

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

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?

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