SEA-PHAGES | All posts created by viknesh

Link to this post \| posted 10 Nov, 2022 16:33
viknesh	plconnerly Vic - we're using all samples with a titer of at least 1 x 10^9. Typically, we can get the minimum 40 ng/ul from a lysate at 5x10^9 pfu/ml, though of course that will vary from phage to phage. Including a ZnCl2 precipitation step has worked well for many, with University of Ottawa now including this as a standard step for all their DNA extractions. It does take a little getting used to, and it is very important to be resuspending the pellet very quickly after the spin, and in EDTA. The precipitation step is harsh and phage DNA is rapidly released after the spin, making them accessible to the nuclease before the denaturant is added. I havent tried this, but perhaps it makes sense to add EDTA to the nuclease-treated lysate before adding ZnCl2. As you troubleshoot, I'd recommend prepping DNA from one lysate both ways (with and without ZnCl2), side by side, to see if the precipitation step works for you. Let me know if you want to chat via Zoom.

Link to this post | posted 10 Nov, 2022 16:33

plconnerly
Vic - we're using all samples with a titer of at least 1 x 10^9.

Typically, we can get the minimum 40 ng/ul from a lysate at 5x10^9 pfu/ml, though of course that will vary from phage to phage. Including a ZnCl2 precipitation step has worked well for many, with University of Ottawa now including this as a standard step for all their DNA extractions. It does take a little getting used to, and it is very important to be resuspending the pellet very quickly after the spin, and in EDTA. The precipitation step is harsh and phage DNA is rapidly released after the spin, making them accessible to the nuclease before the denaturant is added. I havent tried this, but perhaps it makes sense to add EDTA to the nuclease-treated lysate before adding ZnCl2.

As you troubleshoot, I'd recommend prepping DNA from one lysate both ways (with and without ZnCl2), side by side, to see if the precipitation step works for you.

Let me know if you want to chat via Zoom.

Posted in: Phage Discovery/Isolation → DNA Extraction Troubleshooting - Can we skip the nuclease treatment?

Link to this post \| posted 09 Nov, 2022 19:46
viknesh	plconnerly We could use any and all advice about DNA extraction from Gordonia rubripertincta phages. Last year we tried and troubleshot the Wizard DNA Cleanup kit column method with no success. We did phenol:chloroform:isoamyl alcohol as a last resort and managed to get useable DNA from 1 phage. This year we're trying the ZnCl2 method and initial runs have not worked. We've got some troubleshooting planed and one idea we have is to leave out the initial nuclease step. Is that allowed? Have other folks tried that? Can DNA extracted without nuclease treatment be used for sequencing? Thanks! Pam - IU Southeast I'm hoping Dan Russell will chime in here but I believe it is very important to include the nucelase treatment step. Otherwise, the bacterial DNA to phage DNA ratio might be too high, resulting in insufficient sequencing depth for the phage genome. Can I ask about the titer of the samples that are resulting is low DNA extraction?

Link to this post | posted 09 Nov, 2022 19:46

viknesh

plconnerly
We could use any and all advice about DNA extraction from Gordonia rubripertincta phages. Last year we tried and troubleshot the Wizard DNA Cleanup kit column method with no success. We did phenol:chloroform:isoamyl alcohol as a last resort and managed to get useable DNA from 1 phage. This year we're trying the ZnCl2 method and initial runs have not worked. We've got some troubleshooting planed and one idea we have is to leave out the initial nuclease step. Is that allowed? Have other folks tried that? Can DNA extracted without nuclease treatment be used for sequencing?
Thanks!
Pam - IU Southeast

I'm hoping Dan Russell will chime in here but I believe it is very important to include the nucelase treatment step. Otherwise, the bacterial DNA to phage DNA ratio might be too high, resulting in insufficient sequencing depth for the phage genome.

Can I ask about the titer of the samples that are resulting is low DNA extraction?

Posted in: Phage Discovery/Isolation → DNA Extraction Troubleshooting - Can we skip the nuclease treatment?

Link to this post \| posted 21 Sep, 2022 17:09
viknesh	edoddmoh@uottawa.ca Hi! Is there is a reason the smeg top agar is prepared at 2X, and then diluted to 1X? I'm wondering if I can prepare it as a 1L volume, autoclave, and then aliquot into smaller bottles (as with the PYCa top agar). We are phage-hunting with smeg for the first time this semester. Thanks! Liz Hi Liz, Part of the reason top agar is prepared at 2x is so that experiemnts that require the use or large samples of phage (in buffer) do not overly dilute the nutrient in the medium. For our general PHAGES protocols, you should be fine preparing top agar at 1x. 7H9 medium has lots of salts in it, and calcium chloride tends to precipitate out over time. For this reason, calcium chloride is only added before use. Even if you prepare 1x top agar, you might want to only add calcium chloride before use. If you do add it early, please let us know how long it takes for the precipitates to form so that we can share with others. Vic

Link to this post | posted 21 Sep, 2022 17:09

viknesh

edoddmoh@uottawa.ca
Hi! Is there is a reason the smeg top agar is prepared at 2X, and then diluted to 1X? I'm wondering if I can prepare it as a 1L volume, autoclave, and then aliquot into smaller bottles (as with the PYCa top agar). We are phage-hunting with smeg for the first time this semester. Thanks!
Liz

Hi Liz,

Part of the reason top agar is prepared at 2x is so that experiemnts that require the use or large samples of phage (in buffer) do not overly dilute the nutrient in the medium. For our general PHAGES protocols, you should be fine preparing top agar at 1x. 7H9 medium has lots of salts in it, and calcium chloride tends to precipitate out over time. For this reason, calcium chloride is only added before use. Even if you prepare 1x top agar, you might want to only add calcium chloride before use. If you do add it early, please let us know how long it takes for the precipitates to form so that we can share with others.

Vic

Posted in: Phage Discovery/Isolation → 2X smeg top agar

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

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

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?

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